Microfluidic devices for investigating crystallization

ABSTRACT

Microfluidic devices and methods for investigating crystallization and/or for controlling a reaction or a phase transition are disclosed. In one embodiment, the microfluidic device includes a reservoir layer; a membrane disposed on the reservoir layer; a wetting control layer disposed on the membrane; and a storage layer disposed on the wetting control layer, wherein the wetting control layer and the storage layer define a microfluidic channel comprising an upstream portion, a downstream portion, a first fluid path in communication with the upstream and the downstream portions, and a storage well positioned within the first fluid path, wherein the wetting control layer includes a fluid passageway in communication with the storage well and the membrane, and wherein the wetting control layer wets a first fluid introduced into the microfluidic channel, the first fluid comprising a hydrophilic, lipophilic, fluorophilic or gas phase as the continuous phase in the microfluidic channel.

CROSS-REFERENCES TO RELATED APPLICATIONS

This application claims priority from U.S. Patent Application No.62/040,820 filed Aug. 22, 2014.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH

This invention was made with government support under 0754769 awardedfrom the National Science Foundation. The United States government hascertain rights in the invention.

BACKGROUND OF THE INVENTION

1. Field of the Invention

This invention relates to microfluidic devices for investigatingcrystallization.

2. Description of the Related Art

Although protein crystallography can be a very successful technique forstructure determination, membrane proteins continue to presentchallenges to crystallization. It has been reported that two thirds ofpurified proteins fail to produce diffraction quality protein crystals.Of the human membrane proteins, representing one third of the genome,only a few have had their structure solved using X-ray diffraction. Inmany cases, the number of crystallization trials is limited by theavailability of human protein, which does not express well in bacteria,hence the drive to minimize sample volume.

The paradigm guiding many crystallization efforts is that the conditionsfor which an equilibrium crystal phase exists are a small subset among avastly larger set of parameters such as protein concentration, pH,various salts, polymers, temperature, and surfactants. However, it isnot widely appreciated that finding the correct equilibrium conditions,while a necessary condition, is not sufficient to produce crystalsbecause crystallization is a non-equilibrium process. Consequently,crystallization methods that focus on screening large number ofconditions were often incomplete. Additionally, it may be helpful tooptimize the non-equilibrium kinetics of protein crystallization andexploit the crystals that are produced by these methods in order toobtain high quality diffraction data.

Under many previous methods, protein crystals are produced by trial anderror, which necessitates exploring a large number of conditionsconsuming milligrams of protein. Many methods employed in smallnon-automated labs require about 1 microliter of solution per trial.Automation with expensive robotics has lowered volumes to the 100nanoliter (nL) range in some instances. Microfluidic devices can reducethe volume per trial to 1 nL or less in many instances. Such smallvolumes prove useful to screen conditions. However, when crystals areproduced in 1 nL drops, they can be less than 30 microns in diameter,which may be too small for current diffraction methods. Scale-up frommicrofluidic systems also may involve different physics and can bedifficult. Even if large crystals are obtained, then they may berequired to be cryoprotected, which can damage crystals. Finally thecrystals must be aligned in the x-ray beam in many systems.

U.S. Patent Application Publication No. 2012/0190127 to Fraden describesa Crystal Optimizer that is designed to optimize the crystallizationkinetics by systematically varying the kinetic supersaturation profileof the crystallization solution. The technology of U.S. 2012/0190127 canbe used to crystallize proteins on the salvage pathway (promisingcrystals that fail to yield structures), including human membraneG-protein-coupled receptors. Given the paucity of crystallized humanmembrane proteins and the fact that 50% of marketed drugs targetG-protein-coupled receptors, the systems and methods of U.S.2012/0190127 can impact fields such as structural biology andpharmaceutical development.

In the pharmaceutical field, crystal polymorphism can have dramaticdifferences in biological activity between two forms of the same drug.For example, a metastable polymorph may have higher solubility thatleads to an increase in the absorption rate and bioavailability of adrug administered orally. Synthetic and analytic departments of leadingpharmaceutical companies carry out systematic work to detectpolymorphism of their drugs and to find intelligent applications of thisphenomenon. The systems and methods of U.S. 2012/0190127 can benefitsuch systematic work in detecting polymorphism of drugs.

The Crystal Optimizer of U.S. 2012/0190127 addressed the problem ofcrystal creation by determining favorable conditions for crystallizationusing microfluidics. However, there is still a need for furthermicrofluidic technology improvements that allow for a systematic andreversible kinetic control of crystallization trajectory and diffractionstudies of crystallized molecules.

SUMMARY OF THE INVENTION

In one aspect, the present invention provides a microfluidic multiplexdialysis chip for mapping phase diagrams with reconfigurable chemicalpotential.

In another aspect, the present invention provides a technology based onemulsion microfluidics in which drops of protein solution areencapsulated in oil and stabilized by surfactant. We optimize nucleationand growth by generating hundreds of different kinetic pathssimultaneously by varying both temperature and concentration of theprotein solution. Once the optimal kinetic path is determined, weprocess an entire emulsion under optimal conditions to generate onecrystal per drop. The microfluidic device of this version of theinvention can operate with a dialysis membrane, allowing us to optimizekinetic trajectories against various small molecule solutes, such assalts, pH and surfactants. The microfluidic device is compatible for insitu structure studies by X-ray diffraction.

In another aspect, the present invention provides a room temperatureserial crystallography method using a kinetically optimized microfluidicdevice for protein crystallization and on-chip X-ray diffraction. Theemulsion based serial crystallographic technology can use nanolitersized droplets of protein solution encapsulated in oil and stabilized bysurfactant. Once the first crystal in a drop is nucleated, the smallvolume generates a negative feedback mechanism that lowers thesupersaturation, which we exploit to produce one crystal per drop. Wediffract, one crystal at a time, from a series of room temperaturecrystals stored on an X-ray semi-transparent microfluidic chip andobtain a complete data set by merging single diffraction frames takenfrom different unoriented crystals.

In another aspect, the present invention provides devices for supportingcrystals in an X-ray diffraction apparatus and methods for making thedevices.

In another aspect, the present invention provides kits for makingdevices for acquiring X-ray diffraction images of one or more crystals.

It is therefore an advantage of the invention to provide improvedmicrofluidic devices for investigating crystallization.

These and other features, aspects, and advantages of the presentinvention will become better understood upon consideration of thefollowing detailed description, drawings and appended claims.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 shows: in (A), a schematic exploded perspective view of oneembodiment of a microfluidic dialysis Phase Chip according to theinvention, and in (B), a transverse cross-sectional view of a singlestorage well of the Phase Chip in (A).

FIG. 2 shows a transverse schematic overview of fabrication workflow(A1-A3) for the Phase Chip of FIG. 1, with an exploded transverse viewof the final assembly (B) highlighted in the bounding box. Preparationof a poly(dimethylsiloxane) (PDMS) mold is shown in (C). Insertion oftubing directly into the PDMS reservoir is shown in (D). In (E), a topview of final chip assembled with four screws is shown.

FIG. 3 shows in (A), top plan view and perspective view schematics of analternative embodiment of a Phase Chip according to the invention withstorage layers for the single height design (left) and the multipleheight multi-valve design (right), as viewed from top and from the sideperspective view. In (B), there is shown a top view image sequence of anaqueous solution of food dye loaded into the storage layer. Channelcross-sections (height·width) of the device used here are bypass=100μm·100 μm, storage well entry=150 μm·-100 μm, and valves=25 μm·80 μm.The chip was primed with 12 wt % Fluoro-octanol in FC-43, before bluefood dye was injected into the storage wells. The flow rate was 150μL/hr throughout the experiment. White arrows indicate the direction offlow. Scale bar=300 μm. In (C), there is shown, a top view of loadeddevice with all visible 98 wells loaded defect free, illustrating robustsample loading. Scale bar=500 μm.

FIG. 4 shows microfluidic technology according to the invention thatallow for a systematic and reversible kinetic control of crystallizationtrajectory.

FIG. 5 shows: in (A), 30 mg/ml Glucose Isomerase crystallized against aPEG gradient of 20 to 40 wt % PEG of MW 10,000 in 100 mM AmmoniumSulfate, pH 7.3, notice the shallow depth of the crystallization slot,from no crystals (bottom rows), to single crystals, to multiple crystalsper drop to precipitate (top rows); in (B), a close up of inset in (A);in (C), bright field microscopy; and in (D), with Second HarmonicGeneration (SHG) images of Glucose Isomerase crystals in the chip. Thearrow indicates non crystalline protein aggregates that have no SHGsignal.

FIG. 6 shows: in (A), an X-ray transparent chip according to theinvention inside the Cornell University MacChess F1 beamline; in (B), acustom mount is used to hold the thin foil chip in the beam; in (C),Glucose Isomerase crystals inside of the microfluidic device accordingto the invention wherein using a motorized stage, each crystal can becentered in the collimated x-ray beam, and the beam is 100 μm indiameter as indicated by the cross-hair; and in (D), a representativediffraction pattern of a Glucose Isomerase crystal taken at roomtemperature from inside an X-ray chip according to the invention.Crystals diffracted down to 1.37 Å resolution with a mosaicity as low as0.04. The top right quadrant shows the diffraction patterns afterbackground subtraction. A high resolution structure was obtained bymerging data from 72 crystals.

FIG. 7 shows: in (A), optimal crystallization trajectory increasessupersaturation until just one crystal nucleates, then decreasessupersaturation to prevent further nucleation, while remainingsupersaturated enough to promote crystal growth; and in (B), emulsiondroplets with monodisperse crystals were stored in an X-raysemi-transparent microfluidic device according to the invention whereinsequentially collected diffraction frames from multiple individualcrystals were merged to solve the protein structure. The chip could betranslated in x- and y-directions and rotated ±20°.

FIG. 8 shows in A and B, protein concentration as a function of distancefrom a simulation of nucleation and growth in one dimension.Concentration is dimensionless. The dotted line indicates the initialconcentration with a supersaturation of 83.3 at t=0. The sites withconcentrations that exceed the line are in the crystalline phase, whilethose below are in solution. In (A), concentration profile at t=250.Slow nucleation rate of R=1 in dimensionless units. In (B),concentration profile at t=50. Fast nucleation rate of R=27. In (C),average number of crystals per drop as a function of time, (x(t)), fortwo nucleation rates obtained from simulation and fitted to Equation 3,(x(t))=x_(∞)(1−e^(−kt)). Conditions are the same as in (A) and (B). In Dand E, fitting parameters to Equation 3 as a function of drop size fortwo nucleation rates, R=27 and R=1. Arrows indicate size of depletionzone. In (D), the solid lines are the simulated final number of crystalsper drop, x_(∞). Dashed lines are Equation 7, x_(∞)=V=w^(d). In (E), thesolid lines are the simulated rate of crystal formation, k. Dashed linesare Equation 8, k=JV. In (F), a conceptual schematic is shown. A drop ofvolume V can be thought of as x_(∞) smaller, independent drops of volumew^(d)=V/x_(∞).

FIG. 9 shows protein crystallization in emulsion droplets stabilized bysurfactant. Ideal drop sizes were first identified using polydisperseemulsion droplets. Monodisperse emulsion were used to produce identicalcrystals for diffraction experiments. Droplets were stored in arectangular glass capillary. In A to C, polydisperse emulsions of: (A)D1D2 heterodimer from human spliceosomal snRNP particle, (B)concanavelin A, and (C) trypsin. In (D), protein and precipitantsolutions were introduced in a co-flow geometry under laminar flowconditions that prevent mixing upstream of the nozzle where bothsolutions became encapsulated into emulsion droplets. In E and F,monodisperse emulsions of (E) glucose isomerase and (F) lysozymecrystals.

FIG. 10 shows X-ray chip fabrication according to the invention. In thetransverse cross-sectional view of (A), poly(dimethylsiloxane) (PDMS)resin was squeezed into a thin layer onto the SU8-master. After curing,a foil cover was bonded onto the featured PDMS using a silanecoupling-chemistry. Then the reinforced PDMS film was peeled off and thechip was lidded using another foil cover. In (B), top view and (C)cross-section of a device made from cyclic-olefin-copolymer (COC) foilcovers and PDMS. A 5 mm thick slab of PDMS was bonded to the foil coverwith the inlets to form a manifold (see B) for injecting the emulsioninto the chip. FIG. 10D shows a transverse partial cross-sectional viewof a horizontally oriented single layer chip.

FIG. 11 shows: in (A), a monodisperse emulsion being prepared using adedicated dropmaking chip as illustrated in FIG. 9(D) and directlyrouted into the chip for serial crystallography for storage. In (B), weused a laser-cut frame to hold and to port into the X-raysemi-transparent chip. In (C), there is shown an X-ray semi-transparentchip mounted on the goniometer inside the Cornell University CHESS F1beamline. In (D), there is shown glucose isomerase crystals inside ofthe microfluidic device. Using a motorized stage, each crystal can becentered in the collimated X-ray beam. The beam is 100 μm in diameter.In (E), there is shown a representative diffraction pattern of a glucoseisomerase crystal taken at room temperature from inside the chip.Crystals diffracted to 1.4 Å resolution with a mosaicity as low as0.04°. The bottom right quadrant shows the diffraction pattern afterbackground subtraction, using the Adxv diffraction pattern visualizationtool with subtract background option.

FIG. 12 shows part of the final refined structure showing the quality ofthe electron density map. 2Fo-Fc map is in magenta, contoured at 2 Å,Fo-Fc in red (negative) and green (positive), contoured at 3 Å.

FIG. 13 shows cross sectional views of a single membrane differentialpermeation X-ray chip and a double membrane differential permeationX-ray chip according to the invention.

FIG. 14 shows a cross sectional view of an X-ray chip storage containeraccording to the invention.

FIG. 15 shows a schematic of a hydrostatic pressure driven flow systemfor controlling dialysis in an X-ray chip according to the invention.

FIG. 16 shows yeast populations in wells having a volume of 20nanoliters each at a time lapse of approximately one week in a dialysischip according to the invention.

Like reference numerals will be used to refer to like parts from Figureto Figure in the following description of the drawings.

DETAILED DESCRIPTION OF THE INVENTION

In one embodiment, the invention provides a microfluidic deviceincluding a reservoir layer defining a reservoir; a membrane disposed onthe reservoir layer; a wetting control layer disposed on the membrane;and a storage layer disposed on the wetting control layer. The wettingcontrol layer and the storage layer define a microfluidic channelcomprising an upstream portion, a downstream portion, a first fluid pathin fluid communication with the upstream portion and the downstreamportion, and a storage well positioned within the first fluid path. Asused herein, an upstream portion is situated in the opposite directionfrom that in which the fluid flows, whereas a downstream portion issituated in the direction in which the fluid flows. The wetting controllayer includes a fluid passageway in fluid communication with thestorage well and the membrane. The wetting control layer is capable ofwetting a first fluid introduced into the microfluidic channel, thefirst fluid comprising a hydrophilic, lipophilic, fluorophilic or gasphase as the continuous phase in the microfluidic channel.

In certain embodiments of the microfluidic device, the membranecomprises a dialysis membrane, or the membrane comprises a membranepermeable to water, or the membrane comprises a polyethersulfone, or themembrane comprises regenerated cellulose or cellulose ester, or themembrane is hydrophilic.

In certain embodiments of the microfluidic device, the wetting controllayer comprises a fluoropolymer, and the first fluid comprises afluorinated oil. The wetting control layer may comprise a polymericmaterial selected from the group consisting of fluoroakylenes and blendsand copolymers thereof. The wetting control layer may comprisefluorinated ethylene propylene. The storage layer may include afluorophilic coating.

In certain embodiments of the microfluidic device, the reservoir, themembrane, the wetting control layer and the storage layer are reversiblysecured together by clamping or are laminated together. The fluidpassageway may be aligned with the reservoir.

In certain embodiments of the microfluidic device, a plurality ofstorage wells are positioned within the first fluid path, the reservoirlayer defines a plurality of reservoirs, and each reservoir is alignedwith one of the storage wells. The storage layer may comprisepolyurethane, and the reservoir layer may comprise polydimethylsiloxane.The storage layer and the reservoir layer may each comprise plastic,fluoroplastic, or glass.

In another embodiment, the invention provides a microfluidic deviceincluding a microfluidic channel comprising an upstream portion, adownstream portion, a first fluid path in fluid communication with theupstream portion and the downstream portion, a second fluid path influid communication with the upstream portion and the downstreamportion. The second fluid path branches from the upstream portion andreconnects at the downstream portion. A well is positioned within thefirst fluid path, and a plurality of fluid constrictions are in fluidcommunication with the well and the downstream portion. The first fluidpath has less resistance to flow compared to the second fluid path priorto positioning of a first droplet in the well, and the first fluid pathhas greater resistance to flow compared to the second fluid path afterpositioning of the first droplet in the well.

In certain embodiments of the microfluidic device, the well has a wellheight, and each of the fluid constrictions has a constriction heightless than the well height. The well may have a well cross-sectional areameasured perpendicular to a fluid flow direction in the microfluidicchannel greater than a first fluid path cross-sectional area measuredperpendicular to the fluid flow direction in the microfluidic channel.

In another embodiment, the invention provides a device for supportingcrystals in an X-ray diffraction apparatus. The device includes a firstX-ray transparent layer including a microfluidic channel having amicrowell positioned therein; a second X-ray transparent layer includinga reservoir; and a membrane. The first X-ray transparent layer isattached to a first side of a membrane, and the second X-ray transparentlayer is attached to a second opposite side of the membrane such that atleast a portion of an opening of the well and at least a portion of anopening of the reservoir are aligned.

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, a plurality of microwells are positioned withinthe microfluidic channel, the second X-ray transparent layer defines aplurality of reservoirs, and each reservoir is aligned with one of themicrowells. The first X-ray transparent layer may comprise an X-raytransparent material selected from the group consisting of cycloolefinpolymers, cycloolefin copolymers, polyimides, graphene, and siliconnitride, and the second X-ray transparent layer may comprise the X-raytransparent material. The first X-ray transparent layer may comprise acycloolefin copolymer, and the second X-ray transparent layer maycomprise a cycloolefin copolymer. In one embodiment, the first X-raytransparent layer comprises poly(4,4-oxydiphenylene pyromellitimide),and the second X-ray transparent layer comprises poly(4,4-oxydiphenylenepyromellitimide).

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, the first X-ray transparent layer is less than200 microns in thickness, and the second X-ray transparent layer is lessthan 200 microns in thickness. The first X-ray transparent layer may beless than 100 microns in thickness, and the second X-ray transparentlayer may be less than 100 microns in thickness. The first X-raytransparent layer may be less than 50 microns in thickness, and thesecond X-ray transparent layer may be less than 50 microns in thickness.The first X-ray transparent layer may be less than 10 microns inthickness, and the second X-ray transparent layer may be less than 10microns in thickness.

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, the membrane may comprise a dialysis membrane.The membrane may comprise a membrane permeable to water. The membranemay be hydrophilic. The membrane may be less than 50 microns inthickness.

In another embodiment, the invention provides a method for making adevice for supporting crystals in an X-ray diffraction apparatus. Themethod includes the steps of (a) providing a first mold; (b) using thefirst mold to emboss a microfluidic channel in a first X-ray transparentlayer wherein the microfluidic channel has a microwell positionedtherein; (c) providing a second mold; (d) using the second mold toemboss a reservoir in a second X-ray transparent layer; (e) attachingthe first X-ray transparent layer to a first side of a membrane; and (f)attaching the second X-ray transparent layer to a second opposite sideof the membrane such that at least a portion of an opening of the welland at least a portion of an opening of the reservoir are aligned.

In certain embodiments of the method for making a device for supportingcrystals in an X-ray diffraction apparatus, a plurality of microwellsare embossed within the microfluidic channel, a plurality of reservoirsare embossed in the second X-ray transparent layer, and each reservoiris aligned with one of the microwells. The first X-ray transparent layermay comprise an X-ray transparent material selected from the groupconsisting of cycloolefin polymers, cycloolefin copolymers, polyimides,graphene, and silicon nitride, and the second X-ray transparent layermay comprise the X-ray transparent material. The first X-ray transparentlayer may comprise a cycloolefin copolymer, and the second X-raytransparent layer may comprise a cycloolefin copolymer. The first X-raytransparent layer may comprise poly(4,4-oxydiphenylene pyromellitimide),and the second X-ray transparent layer may comprisepoly(4,4-oxydiphenylene pyromellitimide).

In certain embodiments of the method for making a device for supportingcrystals in an X-ray diffraction apparatus, the first X-ray transparentlayer is less than 200 microns in thickness, and the second X-raytransparent layer is less than 200 microns in thickness. The first X-raytransparent layer may be less than 100 microns in thickness, and thesecond X-ray transparent layer may be less than 100 microns inthickness. The first X-ray transparent layer may be less than 50 micronsin thickness, and the second X-ray transparent layer may be less than 50microns in thickness. The first X-ray transparent layer may be less than10 microns in thickness, and the second X-ray transparent layer may beless than 10 microns in thickness.

In certain embodiments of the method for making a device for supportingcrystals in an X-ray diffraction apparatus, the membrane comprises adialysis membrane. The membrane may comprise a membrane permeable towater. The membrane may be hydrophilic. The membrane may be less than 50microns in thickness.

In certain embodiments of the method for making a device for supportingcrystals in an X-ray diffraction apparatus, the method may include thesteps of: (a) introducing a first fluid into the microwell, the firstfluid comprising a protein solution; and (b) introducing a second fluidinto the reservoir, wherein the first fluid, the second fluid and themembrane are chosen such that a crystal forms in the microwell. Themethod may include the step of controlling a temperature of the firstfluid and the second fluid such that the crystal forms in the microwell.

In another embodiment, the invention provides a device for supportingcrystals in an X-ray diffraction apparatus. The device includes a firstX-ray transparent layer; a second X-ray transparent layer; and a storagelayer including a microfluidic channel having a plurality of microwellspositioned therein for containing the crystals. The first X-raytransparent layer is attached to a first side of the storage layer, andthe second X-ray transparent layer is attached to a second opposite sideof the storage layer.

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, the first X-ray transparent layer comprises anX-ray transparent material selected from the group consisting ofcycloolefin polymers, cycloolefin copolymers, polyimides, graphene, andsilicon nitride, and the second X-ray transparent layer comprises theX-ray transparent material. The first X-ray transparent layer maycomprise a cycloolefin copolymer, and the second X-ray transparent layermay comprise a cycloolefin copolymer. The first X-ray transparent layermay comprise poly(4,4-oxydiphenylene pyromellitimide), and the secondX-ray transparent layer may comprise poly(4,4-oxydiphenylenepyromellitimide).

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, the first X-ray transparent layer is less than200 microns in thickness, and the second X-ray transparent layer is lessthan 200 microns in thickness. The first X-ray transparent layer may beless than 100 microns in thickness, and the second X-ray transparentlayer may be less than 100 microns in thickness. The first X-raytransparent layer may be less than 50 microns in thickness, and thesecond X-ray transparent layer may be less than 50 microns in thickness.The first X-ray transparent layer may be less than 10 microns inthickness, and the second X-ray transparent layer may be less than 10microns in thickness.

In another embodiment, the invention provides a method for making adevice for supporting crystals in an X-ray diffraction apparatus. Themethod includes the steps of (a) providing a master mold; (b) using themaster mold to form a storage layer including a microfluidic channelhaving a plurality of microwells; (c) attaching a first X-raytransparent layer to a first side of the storage layer; and (d)attaching a second X-ray transparent layer to a second opposite side ofthe storage layer. The method may include the step of introducing afirst fluid into the microwells, wherein the first fluid comprises aprotein solution or a protein crystal.

In certain embodiments of the method for making a device for supportingcrystals in an X-ray diffraction apparatus, the first X-ray transparentlayer comprises an X-ray transparent material selected from the groupconsisting of cycloolefin polymers, cycloolefin copolymers, polyimides,graphene, and silicon nitride, and the second X-ray transparent layercomprises the X-ray transparent material. The first X-ray transparentlayer may comprise a cycloolefin copolymer, and the second X-raytransparent layer may comprise a cycloolefin copolymer. The first X-raytransparent layer may comprise poly(4,4-oxydiphenylene pyromellitimide),and the second X-ray transparent layer may comprisepoly(4,4-oxydiphenylene pyromellitimide).

In certain embodiments of the method for making a device for supportingcrystals in an X-ray diffraction apparatus, the first X-ray transparentlayer is less than 200 microns in thickness, and the second X-raytransparent layer is less than 200 microns in thickness. The first X-raytransparent layer may be less than 100 microns in thickness, and thesecond X-ray transparent layer may be less than 100 microns inthickness. The first X-ray transparent layer may be less than 50 micronsin thickness, and the second X-ray transparent layer may be less than 50microns in thickness. The first X-ray transparent layer may be less than10 microns in thickness, and the second X-ray transparent layer may beless than 10 microns in thickness.

In another embodiment, the invention provides a device for supportingcrystals in an X-ray diffraction apparatus. The device includes an X-raytransparent layer; and a storage section in the X-ray transparent layer,wherein the X-ray transparent layer and a first side of the storagesection define a microfluidic channel having a plurality of microwellspositioned therein for containing the crystals.

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, the X-ray transparent layer comprises an X-raytransparent material selected from the group consisting of cycloolefinpolymers, cycloolefin copolymers, polyimides, graphene, and siliconnitride. The X-ray transparent layer may comprise a cycloolefincopolymer. The X-ray transparent layer may comprisepoly(4,4-oxydiphenylene pyromellitimide). The X-ray transparent layermay be less than 200 microns in thickness. The X-ray transparent layermay be less than 100 microns in thickness. The X-ray transparent layermay be less than 50 microns in thickness. The X-ray transparent layermay be less than 10 microns in thickness.

In certain embodiments of the device for supporting crystals in an X-raydiffraction apparatus, the microfluidic channel is formed in the storagesection. The device may include a single X-ray transparent layer.

In another embodiment, the invention provides a kit for acquiring X-raydiffraction images of one or more crystals. The kit includes any of thedevices for supporting crystals in an X-ray diffraction apparatus asdescribed above. The kit further includes a crystallization trial devicecomprising: (i) a reservoir layer defining a plurality of reservoirs,(ii) a storage layer defining a microfluidic channel having a pluralityof microwells positioned therein, and (iii) a membrane positionedbetween the reservoir layer and the storage layer, wherein at least aportion of an opening of each microwell is aligned with an opening of areservoir of the reservoir layer.

In another embodiment, the invention provides a kit for acquiring X-raydiffraction images of one or more crystals. The kit includes a firstX-ray transparent layer including a microfluidic channel having aplurality of microwells; a second X-ray transparent layer including oneor more reservoirs; a membrane; and a fastening system for attaching thefirst X-ray transparent layer to a first side of the membrane and forattaching the second X-ray transparent layer to a second opposite sideof the membrane.

In certain embodiments of the kit for acquiring X-ray diffraction imagesof one or more crystals, the first X-ray transparent layer and thesecond X-ray transparent layer are structured such that at least aportion of an opening of each microwell can be aligned with an openingof a reservoir in the second X-ray transparent layer. The first X-raytransparent layer may be less than 100 microns in thickness, and thesecond X-ray transparent layer may be less than 100 microns inthickness. The first X-ray transparent layer may comprise an X-raytransparent material selected from the group consisting of cycloolefinpolymers, cycloolefin copolymers, polyimides, graphene, and siliconnitride, and the second X-ray transparent layer may comprise the X-raytransparent material. The membrane may comprise a dialysis membrane.

The kit may include a crystallization trial device comprising: (i) areservoir layer defining a plurality of reservoirs, (ii) a storage layerdefining a microfluidic channel having a plurality of microwellspositioned therein, and (iii) a membrane positioned between thereservoir layer and the storage layer, wherein at least a portion of anopening of each microwell is aligned with an opening of a reservoir ofthe reservoir layer.

In another embodiment, the invention provides a kit for acquiring X-raydiffraction images of one or more crystals. The kit includes a supply ofX-ray transparent material; a first mold for embossing a microfluidicchannel having a plurality of microwells in a first section of the X-raytransparent material so as to create a first X-ray transparent layer; asecond mold for embossing one or more reservoirs in a second section ofthe X-ray transparent material so as to create a second X-raytransparent layer; a membrane; and a fastening system for attaching thefirst X-ray transparent layer to a first side of the membrane and forattaching the second X-ray transparent layer to a second opposite sideof the membrane.

In certain embodiments of the kit for acquiring X-ray diffraction imagesof one or more crystals, the first mold and the second mold arestructured such that at least a portion of an opening of each microwellis aligned with an opening of a reservoir in the second X-raytransparent layer. The X-ray transparent material may be less than 100microns in thickness. The X-ray transparent material may comprise amaterial selected from the group consisting of cycloolefin polymers,cycloolefin copolymers, polyimides, graphene, and silicon nitride. Themembrane may comprise a dialysis membrane.

The kit may include a crystallization trial device comprising: (i) areservoir layer defining a plurality of reservoirs, (ii) a storage layerdefining a microfluidic channel having a plurality of microwellspositioned therein, and (iii) a membrane positioned between thereservoir layer and the storage layer, wherein at least a portion of anopening of each microwell is aligned with an opening of a reservoir ofthe reservoir layer.

In another embodiment, the invention provides a method for acquiringX-ray diffraction images of crystals. The method includes the steps of:(a) microfluidically producing droplets; (b) feeding the droplets into amicrofluidic channel of an X-ray device wherein the microfluidic channelhas a plurality of microwells positioned therein for containing thedroplets, and the X-ray device is at least partially X-ray transparent;(c) nucleating and growing a crystal in least some of the droplets tocreate a plurality of crystals; and (d) obtaining an X-ray diffractionpattern from the plurality of crystals.

In certain embodiments of the method for acquiring X-ray diffractionimages of crystals, the X-ray device comprises an X-ray transparentlayer attached to a first side of a storage layer, the X-ray transparentlayer and the first side of the storage layer defining the microfluidicchannel. The X-ray device may comprise a first X-ray transparent layer,a second X-ray transparent layer, and a storage layer including themicrofluidic channel, wherein the first X-ray transparent layer isattached to a first side of the storage layer, and the second X-raytransparent layer is attached to a second opposite side of the storagelayer. The droplets may be monodisperse. The droplets may have a sizesuch that a single crystal is grown in each droplet. Each crystal maygrow by permeation of water in the X-ray device. Each crystal may growby dialysis in the X-ray device. Preferably, the method does not includea cryoprotection step.

In another embodiment, the invention provides a device for growingcrystals. The device includes a storage layer having a plurality ofmicrowells positioned therein for containing the crystals; a first X-raytransparent layer attached to a first side of the storage layer; asecond X-ray transparent layer is attached to a second opposite side ofthe storage layer; and a differential permeation membrane attached tothe first X-ray transparent layer.

In certain embodiments of the device for growing crystals, the firstX-ray transparent layer comprises an X-ray transparent material selectedfrom the group consisting of cycloolefin polymers, cycloolefincopolymers, polyimides, graphene, and silicon nitride, and the secondX-ray transparent layer comprises the X-ray transparent material. Thefirst X-ray transparent layer may comprise a cycloolefin copolymer, andthe second X-ray transparent layer may comprise a cycloolefin copolymer.The first X-ray transparent layer may be less than 200 microns inthickness, and the second X-ray transparent layer may be less than 200microns in thickness. The first X-ray transparent layer may be less than100 microns in thickness, and the second X-ray transparent layer may beless than 100 microns in thickness. The first X-ray transparent layermay be less than 50 microns in thickness, and the second X-raytransparent layer may be less than 50 microns in thickness. The firstX-ray transparent layer may be less than 10 microns in thickness, andthe second X-ray transparent layer may be less than 10 microns inthickness.

In certain embodiments of the device for growing crystals, thedifferential permeation membrane may be removably attached to the firstX-ray transparent layer. The differential permeation membrane may have athickness that varies from a first end of the differential permeationmembrane to an opposite second end of the differential permeationmembrane. The differential permeation membrane may be wedge-shaped incross-section. The device may comprise a second differential permeationmembrane attached to the second X-ray transparent layer. The seconddifferential permeation membrane may be removably attached to the secondX-ray transparent layer. The second differential permeation membrane mayhave a thickness that varies from a first end of the second differentialpermeation membrane to an opposite second end of the second differentialpermeation membrane. The second differential permeation membrane may bewedge-shaped in cross-section.

In certain embodiments of the device for growing crystals, a permeationbarrier is attached to the second X-ray transparent layer. At least onefluid port may be in fluid communication with the plurality ofmicrowells. Each fluid port may be in the second X-ray transparentlayer.

In another embodiment, the invention provides a device for containingcrystals. The device comprises a chip and a vial dimensioned to receivethe chip. The chip includes (i) a storage layer including a plurality ofmicrowells positioned therein for containing the crystals, (ii) a firstX-ray transparent layer attached to a first side of the storage layer,and (iii) a second X-ray transparent layer is attached to a secondopposite side of the storage layer.

In certain embodiments of the device for containing crystals, the devicecomprises an aqueous fluid contained in the vial, wherein the aqueousfluid covers the chip when the chip is received in the vial. A seal maycover the aqueous fluid. A source of oil may be contained in the vial,and the source of oil may cover the seal. A conduit may be in fluidcommunication with the source of oil and the storage layer. The vial mayhave an opening such that hydrostatic pressure can push oil into thestorage layer to replace oil that has evaporated. The first X-raytransparent layer and the second X-ray transparent layer may each beless than 100 microns in thickness. The first X-ray transparent layerand the second X-ray transparent layer may each be less than 10 micronsin thickness.

In another embodiment, the invention provides an apparatus forcontrolling a reaction or a phase transition. The apparatus includes amicrofluidic device having a reservoir layer defining a reservoir; adialysis membrane disposed on the reservoir layer; a wetting controllayer disposed on the membrane; and a storage layer disposed on thewetting control layer. The wetting control layer and the storage layerdefine a microfluidic channel comprising a storage well. The wettingcontrol layer includes a fluid passageway in fluid communication withthe storage well and the membrane. The wetting control layer is capableof wetting a first fluid introduced into the microfluidic channel, thefirst fluid comprising a hydrophilic, lipophilic, fluorophilic or gasphase as the continuous phase in the microfluidic channel. The apparatusfurther includes: a source of oil in fluid communication with thestorage well of the microfluidic channel of the microfluidic device; anda source of an aqueous fluid (e.g., a buffer) in fluid communicationwith the reservoir of the microfluidic device. Hydrostatic pressureregulates transport fluxes across the membrane.

In certain embodiments of the apparatus for controlling a reaction or aphase transition, the membrane is a dialysis membrane. In the apparatus,hydrostatic pressure may regulate transport fluxes across the membrane.A pressure controller may regulate transport fluxes across the membrane.The aqueous fluid may be a buffer. In the apparatus, outlets of themicrofluidic channel may be blocked. An outlet of the reservoir may beopen. The apparatus may comprise a microscope for monitoring thereaction or the phase transition in the device. The reaction may be oneor more of the following: steady-state and self-assembly reactions at orfar from equilibrium; perturbation analysis of reaction networks; cellsynchronization; cell and tissue differentiation; and/or chemostatreactions with cells and cell populations. The phase transition may beone or more of the following: crystallization and co-crystallization ofsmall molecules, biological macromolecules, colloids and combinationsthereof; liquid crystal phase transitions; gelation; liquid-liquidseparation; protein folding; and/or DNA melting or condensation. Thereaction may be a chemostat reaction with cells.

In another embodiment, the invention provides an apparatus forcontrolling a reaction or a phase transition. The apparatus includes amicrofluidic device including a microfluidic channel comprising anupstream portion, a downstream portion, a first fluid path in fluidcommunication with the upstream portion and the downstream portion, asecond fluid path in fluid communication with the upstream portion andthe downstream portion. The second fluid path branches from the upstreamportion and reconnects at the downstream portion. A well is positionedwithin the first fluid path, and a plurality of fluid constrictions arein fluid communication with the well and the downstream portion. Thefirst fluid path has less resistance to flow compared to the secondfluid path prior to positioning of a first droplet in the well, and thefirst fluid path has greater resistance to flow compared to the secondfluid path after positioning of the first droplet in the well. Incertain embodiments of the microfluidic device, the well has a wellheight, and each of the fluid constrictions has a constriction heightless than the well height. The well may have a well cross-sectional areameasured perpendicular to a fluid flow direction in the microfluidicchannel greater than a first fluid path cross-sectional area measuredperpendicular to the fluid flow direction in the microfluidic channel.

The reaction may be one or more of the following: steady-state andself-assembly reactions at or far from equilibrium; perturbationanalysis of reaction networks; cell synchronization; cell and tissuedifferentiation; and/or chemostat reactions with cells and cellpopulations. The phase transition may be one or more of the following:crystallization and co-crystallization of small molecules, biologicalmacromolecules, colloids and combinations thereof; liquid crystal phasetransitions; gelation; liquid-liquid separation; protein folding; and/orDNA melting or condensation. The reaction may be a chemostat reactionwith cells.

Within this specification, embodiments have been described in a waywhich enables a clear and concise specification to be written, but it isintended and will be appreciated that embodiments may be variouslycombined or separated without parting from the invention. For example,it will be appreciated that all features described herein are applicableto all aspects of the invention described herein.

EXAMPLES

The following Examples are provided in order to demonstrate and furtherillustrate certain embodiments and aspects of the present invention andare not to be construed as limiting the scope of the invention.

Example 1

In this example, we demonstrate a microfluidic multiplex dialysis chipfor mapping phase diagrams with reconfigurable chemical potential.

Overview

The Phase Chip described in Shim et al., “Using Microfluidics toDecouple Nucleation and Growth of Protein Crystals”, Crystal Growth &Design 2007, Vol. 7, No. 11, pages 2192-2194, compartmentalizes proteincrystallization trials into nanoliter sized emulsion droplets and canscreen up to several thousand kinetic crystallization pathways inparallel, while consuming nanoliter, or even picoliter amounts persample well. In this example, we introduce a new Phase Chip design thatutilizes a dialysis membrane, which greatly extends the range ofapplications, as any solute smaller than the molecular weight cut-off ofthe membrane can be dialyzed into and out of the sample wells. The chipis operated by controlling osmotic and hydrostatic pressure to regulatetransport fluxes across the membrane. Because of its modular design, thechip can be reused multiple times and also harvest crystals from thechip for structure determination by x-ray diffraction.

Introduction

Microfluidic technology allows for exceptional control of solutionconditions in space and time, which has been exploited to map phasediagrams. In particular, crystallization in microfluidic devices hasbeen investigated. In all these microfluidic approaches tocrystallization and in particular with respect to proteincrystallization however, it was neglected that crystal nucleation andcrystal growth require opposing degrees of supersaturation. To decouplenucleation and growth by means of controlling the chemical potential inthe crystallization trial, a Phase Chip as described in Shim et al. hasbeen developed. While the Phase Chip technology of Shim et al. hassuccessfully been demonstrated, its permeation design is limited incertain ways. In the Shim et al. device, the storage layer and thegradient layer are built from separate PDMS pieces, which are covalentlybonded together. Thus, chips are single use only. Because chemicalcoupling takes place across a PDMS membrane in the Shim et al. device,the flux between storage and gradient layer is limited. Small non-polarmolecules permeate quickly, but water permeates very slowly and chargedmolecules do not permeate at experimentally relevant timescales. Thesepoor transport characteristics favor very thin PDMS membranes that arevery fragile and often rupture causing failure of the chip. To overcomethese limitations, we replaced the PDMS membrane with a regeneratedcellulose dialysis membrane (see FIG. 1). Here solute exchange islimited only by the molecular cut-off level of the chosen membrane, andions, acids, bases or bigger molecules such as pH-buffers or evenpolymers can diffuse across the membrane. The dialysis Phase Chip 20 ofFIG. 1 comprises two microfluidic layers 21, 22 that are separated by asemipermeable dialysis membrane 23 and a perforated Teflon®polytetrafluoroethylene foil 24 for wetting control. The storage layer21 on top is a matrix of a few hundred and up to a few thousand storagewells 25 that each can hold an isolated protein solution sample. Thesample droplets 27 can interact osmotically through the dialysismembrane 23 with the solution perfused in the PDMS reservoir layer 22located at the bottom of the chip. Water and solutes can exchangebetween sample droplet and reservoir 28 across the membrane.

Because of the continuous standing column of water in a dialysismembrane, pressure gradients across the membrane can equilibrate throughreverse osmosis. This makes it difficult to dialyze nanoliter volumes inparallel. In the microfluidic device of this example, we can exploitosmosis and reverse osmosis to continuously and reversibly re-formulateeach crystallization cocktail. We can also decouple protein crystalnucleation and growth, by first quenching into a deep supersaturationand then quench back to a low supersaturation. In one non-limitingexample embodiment, the chip is a clamped assembly, where the storagelayer 21 is made from polyurethane and the reservoir layer 22 from PDMS.The flexible PDMS reservoir is mechanically supported by an acrylic(e.g., poly(methyl methacrylate)) bottom piece 26 to provide a goodseal. Because of this modular design where both layers 21, 22 areclamped together, rather than covalently combined, chips can be reusedmultiple times. However, it is also contemplated that the storage layer21 and the reservoir layer 22 can be all be fabricated from plastics,fluoroplastics, and/or glass.

Device Fabrication and Assembly

The dialysis Phase Chip of this example was built by combiningsoft-lithography and replica molding with custom laser cut parts. FIG. 2shows a schematic overview of fabrication workflow (A1-A3) for thedialysis Phase Chip with the final assembly (B) highlighted in thebounding box. In the first step, we prepared a PDMS mold 31 (A1 and C)so that we could later cast the polyurethane storage layer 21 (A2). Wedirectly casted the tubing for interfacing with the storage layer intothe polyurethane. The dangling end was trimmed later. The reservoir wasdirectly molded from a SU8 master (A3). Here we inserted tubing 36directly into the PDMS reservoir (D) and wound the tubing through theacrylic back. In (E), a top view of final chip 20 assembled with fourscrews is shown. Scale bars are 1 cm.

Specifically, for fabricating the storage layer, we manufactured an‘inverse’ negative resist master where the features are wells surroundedby higher SU8 resist 33. We mounted this wafer 34 into an acryliccasting frame 35 (see FIGS. 2 A1 and C) to cast a PDMS replica that canact as a mold for the polyurethane resin (Crystal Clear 204, Smooth-On,Inc.), which after curing formed the storage layer lid 21. For easychip-to-world interfacing, we inserted 300 μm inner diameter polyetherether ketone tubing 36 (PEEK tubing, from Cole Parmer) into the PDMSmold, so that it became embedded into the polyurethane piece (see FIG.2, A2). For this we punched through holes into the PDMS mold using a0.75 mm biopsy punch (Uni Core, Harrison) and inserted the tubing 36into these holes to seal off and prevent resin from flowing into thetubing. A small knot in the tubing 36 helped to firmly embed it into thestorage lid 21. The PDMS mold was then degassed in a desiccator for 30minutes, before the polyurethane resin was cast into the mold. We thenfurther degassed until all air-bubbles trapped in the resin had moved tothe surface. Usually this was after about 1 hour. We then opened thedesiccator and gently popped remaining bubbles using a Pasteur pipette,or a stream of compressed air. We let the polyurethane cure in the ovenat 80° C. overnight. After removing the cured polyurethane top from thecast, we drilled through holes and trimmed the dangling end of thetubing. After a quick clean of the device with compressed air, we plasmaactivated the polyurethane at 500 mTorr (±50 mTorr) O₂ plasma at 60 wattfor 1 minute, before dip-coating the lid in a 1:50 dilution of Cytop 109AE in CTsolv 100E (both Bellex International). Cytop is a fluorophiliccoating that prevents protein unfolding on the channel surface. To fullycross-link the Cytop to the polyurethane surface, we incubated the lidin the oven at 100° C. over night.

The wetting control layer 24 was cut from a 50 μm thick fluorinatedethylene propylene (FEP) foil (McMaster Carr) using a VLS3.50 Versalaser cutter with 50 watt Imaging Cartridge with High Power DensityFocusing Optics (HPDFO). (FIG. 2(B) shows an alternative Teflon® foilfor the wetting control layer.)

The PDMS reservoir 22 was cast on a traditional SU8 master, wherefeatures built up as posts defined the channels in the PDMS piece (seeFIG. 2 A2). To interface into the PDMS reservoir, we punchedthrough-holes using a 0.75 mm biopsy punch (Uni Core, Harrison) anddirectly inserted the tubing into the holes (see FIG. 2 B). The 73 μmflat sheet regenerated cellulose dialysis membrane with 6000 molecularweight cut-off (Bel-Art Products, Peaquannock, N.J.) was incubated for15 minutes in ultrafiltered water (Millipore Elix 3) to wash awayglycerol and other additives used for storing the membrane. We thenwashed again for 15 minutes in 10 mM EDTA to remove residual metal ions.

The acrylic back 26 was cut to shape using a VLS3.50 Versa laser cutterwith 50 watt Imaging Cartridge with HPDFO and then manually threaded sothat four screws could pull the chip-sandwich together.

To assemble the chip (see FIG. 2 E), we positioned the interfaced PDMSreservoir layer 22 on the acrylic back 26. The rinsed dialysis membrane23 of desired pore-size was then gently dried using Kimwipe tissues suchthat the membrane was moist inside, but no water puddles remained on itssurface. We dispensed 100 μl FC-43 (Sigma Aldrich) onto the dialysismembrane before depositing the wetting control layer 24 on top. Another100 μl FC-43 were dispensed onto the FEP foil 24 before placing thepolyurethane storage layer 21 on top. The FC-43 oil lubricated the FEPfoil, so that it could be easily aligned later. We inserted the screwsto clamp the sandwich together, aligned the wetting control layer 24with the polyurethane storage lid 21 and finally tightened the screwsuntil the features in the PDMS reservoir layer 22 were barely beginningto distort from the pressure. We finally primed the chip by dead-endfilling via tubing 38 the aqueous reservoir solution into the reservoir28 and 12 wt % Fluorooctanol in FC-43 39 into the storage layer 21 untilno air-bubbles remained trapped in the chip. After each use, allcomponents were washed in a sonicator bath with 1 vol % Helmanex and 1wt % Zonyl® FSN-100 fluorosurfactant for 15 minutes, then repeatedlyrinsed with ultrafiltered water, dried with compressed air and storedaway for reuse.

Loss-Free Sample Loading Using Capillary Valving

We stored the sample in cylindrical wells. All storage wells wereconnected in series by a continuous serpentine channel through which onewell was loaded after the other (see FIG. 3). To eliminate sample lossfrom channel dead-volume, we exploited the capillary valve based“store-then-create” loading technique. In brief, the chip was primedwith a fluorinated oil before the aqueous sample was injected into thedevice. The surface tension at the oil-water interface between primingoil and aqueous sample resulted in a pressure difference across theinterface. This Laplace pressure can be calculated by the Young-Laplaceequation as

${\Delta \; P} = {\in \left( {\frac{1}{Rx} + \frac{1}{Ry}} \right)}$

with R_(x) and R_(y) as the main radii of curvature and E being thesurface tension of the interface. To minimize its energy, the interfacehas to minimize its surface which is equivalent to maximizing its mainradii of curvature at constant volume. A low curvature interface in awide channel has a lower Laplace pressure then a high curvatureinterface in a narrow channel segment. Therefore, the sample plugpreferentially entered and flowed through the wide bypass channelinstead of flowing through the narrow capillary valve channel (see FIG.3). The sample plug was then followed again by oil, which separated thesample in the wells into independent droplets.

Compared with the previous design of Shim et al., we improved the PhaseChip storage layout in several ways (see FIG. 3 A). First, we routed thebypass closer around the wells to condense the design. By this weincreased the well density by 27%. Second, we made the chipmanufacturing more robust by replacing the fragile thin and tall valvein the single height design with a shallow and wide valve in a multiheight design. This greatly improves master durability and also allowedus to cast plastic chips from silicone rubber molds. Third, we improvedloading efficiency by introducing multiple parallel valves. This, inanalogy to electric resistors connected in parallel, reduced thehydrodynamic resistance of the valve section and allowed us to load thechip reliably with flow rates of up to 150 μl/hr, which was a more thana 3 fold increase over the single valve design.

Looking at FIG. 3, a microfluidic channel 40 includes an upstreamportion 41, a downstream portion 42, a first fluid path 43 in fluidcommunication with the upstream portion 41 and the downstream portion42, a second fluid path 44 in fluid communication with the upstreamportion 41 and the downstream portion 42. The second fluid path 44branches from the upstream portion 41 and reconnects at the downstreamportion 42. A well 45 is positioned within the first fluid path, and aplurality of fluid constrictions (valves) 46 are in fluid communicationwith the well 45 and the downstream portion 42. The first fluid path hasless resistance to flow compared to the second fluid path prior topositioning of a first droplet in the well 45, and the first fluid pathhas greater resistance to flow compared to the second fluid path afterpositioning of the first droplet in the well 45. The well 45 has a wellheight, and each of the fluid constrictions 46 has a constriction heightless than the well height. The height of each of the fluid constrictions46 may be the same or different from the other fluid constrictions 46.The well 45 may have a well cross-sectional area measured perpendicularto a fluid flow direction D in the microfluidic channel greater than afirst fluid path cross-sectional area measured perpendicular to thefluid flow direction D in the microfluidic channel 40. The first fluidpath 43, the second fluid path 44, the well 45, and the plurality offluid constrictions (valves) 46 may be arrayed in a repeatingsymmetrical manner as shown in FIGS. 3 B and 3 C.

Assessing Crystal Quality

Crystal nucleation is a non-equilibrium, dynamic process and timelydetection of crystal nuclei allows prompt optimization ofcrystallization recipes. To finely titrate our dialysis crystallizationtrials into the crystallization zone, we needed to be able to detect thesmallest possible nuclei, or ideally even sub-critical nuclei, whichhave not yet grown larger then the critical size associated with thenucleation barrier. To identify quench profiles that yield a crystallinephase instead of a kinetically arrested gel, we used Second HarmonicGeneration imaging. Crystals grown in the chip were then harvested tocollect X-ray diffraction data.

SHG Imaging

Second Harmonic Generation (SHG) is the emission of radiated, coherentlight at exactly twice the frequency of the incident light.Non-centrosymmetric molecular polarizability can lead to SHG and thusany chiral protein crystal can give a SHG signal, while disordered orcentrosymmetric packings of the same individual proteins cannot. Thesedifferent susceptibilities, make it a powerful detection technique forprotein crystals, as even microcrystals can be selectively imagedagainst a background of solvated protein or amorphous proteinaggregates. Exploiting the ratio of the forward-to-backward detectedSHG, one can detect sizes of green fluorescent protein microcrystallitesand derive a general theoretical detection limit for proteincrystallites of down to 100 nanometers in diameter under lowmagnification with 10× objective. These are detection limits not rivaledby traditional fluorescent, or polarization microscopy methods. Also,usually protein aggregates in solution produce substantial backgroundfluorescence, but no detectable SHG. Similarly, salt crystals arebirefringent too, but do not show SHG.

SHG is a scattering process, so there is no bleaching and because it isa 2-photon effect, there is no background SHG. The SHG signal from aprotein arises primarily from the amide transition of amino acidresidues. Summing the individual hyperpolarizability terms over allresidues yields the SHG susceptibility tensor of a single protein. Fromthe SHG susceptibility of a single protein, the SHG signal from aprotein cluster can be calculated.

Glucose isomerase crystallizes with orthorhombic symmetry in the spacegroup I222. It is known to have a good SHG signal. We grew crystals byloading 30 mg/ml glucose isomerase at 20 wt % PEG 10000, 100 mM ammoniumsulfate, pH 7.3 and subsequently quenching the whole microfluidic chipof this example to 30 wt % PEG 10000, 100 mM ammonium sulfate, pH 7.3.We then sealed all outlets and incubated the chip in a water bath at 4°C. for several days. Crystals were observed inside the chip using brightfield microscopy and SHG imaging, using the SONICC imaging platform(Formulatrix, Waltham, Mass., USA) with 10× objective. Glucose isomerasecrystals gave strong SHG signal, while no SHG signal was observed fromamorphous aggregation found in a few wells. We did not detectsignificant background from the chip in SHG. We concluded that the chipwith its different components is well suited for SHG imaging.

Harvesting Crystals for X-Ray Crystallography

To mount crystals for cryo crystallography, we carefully opened the chipof this example, by gently pressing the lid down while removing all fourscrews. We then took the storage lid 21 off the chip and applied 1 ml 40wt % PEG 10000, 100 mM ammonium sulfate, pH 7.3 onto the lid and alsoonto the polytetrafluoroethylene foil 24 and dialysis membrane 23 leftbehind on top of the reservoir layer 22, as crystals stayed on bothhalves. The 40 wt % PEG buffer was used as a cryo protectant and also tokeep the crystals moist for the duration of the looping.

In our FC-43 oil with 12 wt % fluorooctanol system, emulsion dropletswere not stable against coalescence. Opening the chip and deposition ofnew buffer disrupted the emulsion stored in the chip. We found somecrystals to remain in the stored wells, or attached to thepolytetrafluoroethylene foil 24, while other crystals were freelyfloating in the puddle of cryo-protectant covering the chip.

We looped crystals using standard Nylon loops (Hampton Research). Loopedcrystals where immediately cryo frozen by plunging into liquid nitrogen.Crystals remained stored in liquid nitrogen until X-ray diffraction datawas collected at the MacChess F1 beam line at Cornell University in acryostream (N₂(g)). We took 40 consecutive frames with 1° rotation and 1second exposure for a total of 46 crystals. Most crystals diffracted tobetter then 1.5 Å resolution. We defined the resolution cut-off to wherethe Bragg peak intensity dropped below twice the background intensity.The mosaicities for the crystals in our data set fell into a range of0.2 and 0.4. The best crystal in the set had a diffraction better thenthe edge of the detector at 1.17 Å with mosaicity of 0.15 to 0.22.

Even though our approach to open the dialysis Phase Chip resulted inmechanical disruption of the emulsion droplets and hence the crystals inthem, we could consistently loop high-quality crystals for X-raycrystallography from the chip. To minimize mechanical disruption whenopening the chip, the polytetrafluoroethylene sheet 24 for wettingcontrol could be covalently attached to the storage lid 21. This wouldensure that droplets would stay intact when retrieving the storage lidfrom the chip. Each droplet could then be accessed independently.

Conclusions

In this example, we designed a new microfluidic dialysis chip, tokinetically probe phase diagrams in a high throughput manner. Exploitingosmosis and reverse osmosis, we performed proof of principle experimentscrystallizing glucose isomerase. We confirm that protein crystallizationcan be monitored using Second Harmonic Generation and that crystals canbe harvested from the chip to collect high resolution X-ray diffractiondata. We envision extending the capabilities of the reservoir layer toformulate spatial concentration gradients along one or two dimensions,or to include formulator capabilities. Ultimately we envision applyingthe dialysis Phase Chip of this example to optimize membrane proteincrystallization trials with respect to optimal detergent concentration,which can not be accomplished in classic crystallization trials. We alsoenvision a dialysis chip compatible with polarization microscopy to beable to investigate assembly and disassembly of biological hydrogelssuch as intermediate filament assemblies or amyloid fibrils.

Thus, we improved the permeation Phase Chip described in Shim et al.,“Using Microfluidics to Decouple Nucleation and Growth of ProteinCrystals”, Crystal Growth & Design 2007, Vol. 7, No. 11, pages2192-2194, such that the composition of the dialysis membrane can bechosen arbitrarily. The example design of FIG. 1 may include fourseparate layers in order from top to bottom: (1) a storage layer 21; (2)a wetting control layer 24 (e.g., FEP); (3) a membrane 23; and (4) achemical potential reservoir 22. A protein solution 27 can be injectedin the storage layer 21. The wetting control layer 24 (e.g., FEP)ensures that all walls of the storage loading channels are wetted by thefluorinated oil 39. The wetting control layer 24 (e.g., FEP) maycomprise a fluorinated sheet with one perforation at the location ofeach storage chamber in the first layer. Below the wetting control layer(e.g., FEP) is a dialysis membrane 23. This example dialysis membrane ofthe invention enables the programmed exchange of pH, salts, surfactants.and other small molecule solutes into and out of the storage layer. Thisfeature greatly extends the range of potential applications for thedialysis chip as it now becomes feasible to conduct cell cultureexperiments on chip, but also phase diagrams at constant ionic strengthcan be recorded. We simplified chip manufacturing by adopting a modulardesign. The device is assembled by clamping together the four pieces.Previously, devices were irreversibly bonded together into a singlepiece. However, it is also contemplated that the reservoir 22, themembrane 23, the wetting control layer 24 and the storage layer 21 maybe laminated together.

The dialysis membrane of the invention has applications in the screeningand optimization of kinetic trajectories (e.g., protein crystallizationconditions, stem cell differentiation pathways, etc.) and in the mappingof phase diagrams (e.g., colloidal systems, cell viability assays,chemical reaction kinetics, etc.). The dialysis membrane of theinvention has advantages including: (1) it is a 100% reusable chip; (2)it is modular such that any kind storage, reservoir and membrane can becombined; (3) samples can be retrieved by easily disassembling thedevice enabling easy access to the processed samples (e.g., crystalslooped. cells harvested); and (4) easy fabrication (embossing with nolidding needed).

Example 2

In this example, we demonstrate microfluidic devices to map proteinphase diagrams and nucleation kinetics for in situ x-ray diffraction ofprotein crystals.

Overview

We developed a technology based on emulsion microfluidics in which 1nanoliter drops of protein solution are encapsulated in oil andstabilized by surfactant. Crystallization is a stochastic process; wedetermine nucleation kinetics by measuring thousands of identical drops.We optimize nucleation and growth by generating hundreds of differentkinetic paths simultaneously by varying both temperature andconcentration of the protein solution. Once the optimal kinetic path isdetermined, we process an entire emulsion under optimal conditions togenerate one crystal per drop. We improved the Phase Chip microfluidicdevice described at Shim et al., “Using Microfluidics to DecoupleNucleation and Growth of Protein Crystals”, Crystal Growth & Design2007, Vol. 7, No. 11, pages 2192-2194. For example, the microfluidicdevice of this example can operate with a dialysis membrane, allowing usto optimize kinetic trajectories against various small molecule solutes,such as salts, pH and surfactants. Also, the microfluidic device of thisexample is compatible for in situ structure studies by synchrotrondiffraction.

Introduction

There is no guarantee that a given protein has a crystalline phase, buteven existence of an equilibrium phase is not sufficient for a crystalto form because the transformation of a protein solution to a crystal isgoverned by two nonequilibrium processes: nucleation and growth.Consequently, supersaturation kinetics play an essential role incrystallization and we believe that the optimal crystallization strategyshould screen kinetic trajectories involving variables such as depth ofsupersaturation, duration of supersaturation, and sample volume. Whileother microfluidic crystallization platforms capable of in situdiffraction have been developed, to our knowledge no technique availableallows for a systematic and reversible kinetic control of thecrystallization trajectory. This entails finding conditions on-chip forwhich one crystal is grown per drop and then isolating hundreds of dropsstored on a x-ray transparent microfluidic chip. Single,non-cryoprotected crystals are too small to collect a completediffraction set, but a full data set can be obtained by combining manysingle diffraction patterns.

FIG. 4 shows in (A), ideal crystallization trajectory increasessupersaturation until just one crystal nucleates, then decreasessupersaturation to prevent further nucleation, but promote crystalgrowth. In FIG. 4 (B), the stored protein droplet is osmotically coupledto a chemical potential reservoir using a semipermeable membrane. We useeither a dialysis membrane to exchange small molecule solutes and salt,or an elastomeric membrane, to only permeate water and non-polar solutesbetween both microfluidic layers. With a dedicated sample stage we cansimultaneously quench concentration and generate temperature gradientsto rapidly screen hundreds to thousands of different kineticcrystallization trajectories in parallel. In FIG. 4 (C), using Lysozymeas a model protein, we find sharp transitions from non-crystallizing tocrystallizing trajectories. In FIG. 4 (D), there is shown a schematictop-view of our set-up, illustrating the perpendicular arrangement oftemperature and concentration control. Both can be controlledreversibly, allowing us to decouple protein crystal nucleation andgrowth.

Experimental

The dialysis microfluidic device of this example is built combiningstandard soft-lithography and replica molding with custom laser cutparts. For fabricating the storage layer, we manufacture an ‘inverse’negative resist master where the features are wells surrounded by higherSU8 resist. We mount the wafer into an acrylic casting frame to cast aPDMS replica that can act as a mold for the polyurethane resin (CrystalClear 204, Smooth-On, Inc.), which once cured forms the storage layerlid. The PDMS reservoir is cast on a traditional SU8 master, wherefeatures built up as posts define the channels in the PDMS piece. Thechip is then assembled by placing the dialysis membrane between thestorage and reservoir layers and clamped together with the help of amatching acrylic back that was cut to shape with a laser cutter.

The X-ray Phase Chip microfluidic device of this example is fabricatedby bonding 50 μm thin cyclic-olefin-copolymer (COC) foil (TOPAS®) ontothin PDMS slabs that were fabricated by spin-coating PDMS onto a master.We then peel the storage and the reservoir layer with the COC lids fromthe master, punch through holes and bond them to a prefabricated ˜40 μmthin PDMS membrane such that the PDMS membrane lids both halves of thechip. The X-ray Phase Chip is then mounted into a dedicated acrylicframe with fluid connectors to interface the chip.

Results and Discussion

As a proof of principle experiment, we crystallized Glucose Isomerase(Hampton Research) in a PEG gradient (see FIG. 5). We optimize kineticcrystallization trajectories using Second Harmonic Generation (SHG)imaging to distinguish amorphous protein aggregates from small crystalclusters (FIG. 5 C&D). To directly collect diffraction data fromcrystals grown in chip, we use PDMS-COC hybrid devices that wereoptimized for low X-ray background (FIG. 6). By encapsulating thecrystallization cocktail into emulsion droplets (FIG. 6 C), we yieldhighly monodisperse crystals, with one crystal per drop. We merged 230x-ray diffraction image frames taken at room temperature, with onedegree rotation each from a total of 70 Glucose Isomerase crystals intoa single dataset and solved the structure by molecular replacement downto 1.9 Å resolution (R=0.1579). We took about 5 diffraction frames percrystal with exposure times of 1 second each. Because the crystals werenot cryo-frozen, considerable radiation damage was accrued and alldiffraction peaks had vanished after 10 to 15 seconds accumulatedexposure. Ideally, a dataset merged from more crystals with only singlesub-second exposures each would yield a better resolution structure.

Conclusion

As demonstrated in this example, we developed new microfluidic tools tosupport the protein crystallography community. By incorporating adialysis membrane into a Phase Chip microfluidic device, we can screenhundreds to thousands of different crystallization conditions withcomplex kinetic trajectories in parallel. Thus, we can optimizecrystallization recipes to grow monodisperse crystals, with one crystalper drop. These crystal emulsions can then be transferred to the X-raytransparent chip, to collect room temperature diffraction data from manyidentical crystals. The dialysis Phase Chip microfluidic device of thisexample enables new avenues for single cell and small populationchemostat experiments, or multiplex perturbation reactors to mapnonlinear chemical kinetics of complex reaction networks as found inmany biochemical pathways.

Example 3

In this example, we demonstrate room temperature serial crystallographyusing a kinetically optimized microfluidic device for proteincrystallization and on-chip X-ray diffraction.

Overview

In this example, we demonstrate that we have developed an emulsion basedserial crystallographic technology in which nanoliter sized droplets ofprotein solution are encapsulated in oil and stabilized by surfactant.Once the first crystal in a drop is nucleated, the small volumegenerates a negative feedback mechanism that lowers the supersaturation,which we exploit to produce one crystal per drop. We diffract, onecrystal at a time, from a series of room temperature crystals stored onan X-ray semi-transparent microfluidic chip and obtain a complete dataset by merging single diffraction frames taken from different unorientedcrystals. As proof-of-concept, we solved the structure of glucoseisomerase to 2.1 Å, demonstrating the feasibility of high-throughputserial X-ray crystallography using synchrotron radiation.

1. Introduction

In conventional protein X-ray crystallography, a complete data set isideally obtained from a single crystal, which typically requires arelatively large crystal that has been successfully cryocooled. Serialcrystallography takes the opposite approach: complete diffraction setsare assembled from a large number of individual diffraction framesacquired from small, single, unoriented crystals that are notcryoprotected. Complete coverage of the Ewald sphere is obtained byassembling individual diffraction frames into a single data set. Theideal crystals for serial crystallography are large enough andsufficiently defect free to diffract to high resolution, are produced inlarge quantity, and are sufficiently identical to facilitate merging ofdiffraction frames.

Serial crystallography with non-cryocooled crystals has severaltechnical advantages over conventional methods. First, the crystals canbe small, which increases the potential for growing crystals in thefirst place. Second, it avoids the roughly ten-fold increase in crystalmosaicity typically encountered during cryoprotection and eliminates theneed to search for cryoprotectant conditions. Although non-cryoprotectedcrystals suffer radiation damage at a roughly hundred times higher ratethan cryoprotected crystals, there is little disadvantage associatedwith using many non-cryocooled crystals to obtain a complete data set ifthe crystals are easy to produce, plentiful, and easy to handle.

The ideal crystallization procedure, illustrated in FIG. 7(A), toproduce protein crystals for any form of crystallography, includingserial crystallography, comprises slowly increasing the supersaturationof a protein solution until the moment that a single crystal isnucleated. Then, once the first nucleation event occurs, thesupersaturation is reduced enough to prevent further nucleation, whileremaining supersaturated enough to grow the crystal. Ideally, the growthconditions should be slow enough to allow for annealing of defects, andthe procedure must be capable of producing crystals in large numbers andof identical size. Additionally, the technology to produce crystals mustbe simple and inexpensive if serial crystallography is to become adoptedby the structural biology community.

The challenge is to design such a method. The well knownCounter-Diffusion method produces a series of kinetic supersaturationprofiles that rise and fall as illustrated in FIG. 7(A). However, boththe time at which the supersaturation maximum occurs and the value ofthe supersaturation maximum are independent of the nucleation event. Themaximum supersaturation varies along the capillary length and with along capillary the chances are improved that somewhere along thecapillary there will be a location where the maximum supersaturationwill coincide with the first nucleation event. However, this methodrequires long capillaries and is not optimal for volumes under 1nanoliter. Furthermore, Counter-Diffusion requires that the precipitantand protein have greatly different diffusion constants, so it issuitable for low molecular weight precipitants, such as salt, but notfor macromolecules, such as poly(ethylene glycol) (PEG).

Another issue complicating design of the ideal profile of FIG. 7(A), isthat at constant supersaturation nucleation is a random process,rendering it impossible to a priori know when to decreasesupersaturation, which should coincide with the first nucleation event.One way to generate the ideal supersaturation profile would be tomonitor the supersaturated solution with a technique, such as SecondHarmonic Generation (SHG) microscopy, that is sensitive to the formationof small crystals and then, once the first crystal is detected, lowerthe supersaturation. However, this scheme will be cumbersome toimplement in the high throughput case of processing hundreds tothousands of samples. An alternative method is desired.

Microfluidically produced, monodisperse, emulsions have previously beenused to produce drops of supersaturated protein solution in which eachdrop nucleates a single crystal. This situation is ideal for serialcrystallography for a number of reasons. Since only one crystalnucleates per drop, all the supersaturated protein in solution isdelivered to a single crystal, making that crystal as large as possible.Microfluidic precision allows preparation of emulsion droplets withvariations in size of a few percent only, even at high flow rates.Furthermore, because of the small length scales in microfluidics,convection is suppressed and flows are laminar. Taken together,processing proteins using microfluidics leads to crystals of the uniformsize that are grown under identical conditions, which has the effect ofcreating crystals that have similar characteristics, such as unit celland degree of disorder. Having identical crystals facilitates merging ofdiffraction data sets taken from different crystals.

In the microfluidic device described in this example, we first producedrops containing protein. Then, exploiting surface tension forces, weguide drops to 8,000 storage sites on-chip. Next, we increasesupersaturation to induce crystallization in such a way as to produceone crystal per drop. Finally, we sequentially collect diffraction fromindividual crystals and merge data sets in order to solve the proteinstructure (see FIG. 7B).

Producing and diffracting from crystals in the same device eliminatesthe laborious and potentially damaging steps of looping and extractingthe crystal from the mother liquor. Various microfluidic crystallizationplatforms compatible with in situ diffraction have been developed.However, these devices incorporated valves in the chip, thus renderingthem expensive to manufacture and difficult to operate. Othertechnologies are low-throughput, or need a second round of scale-up tolarger capillaries to produce crystals large enough to collectdiffraction data.

2. One Crystal Per Drop Through Compartmentalization

The production of one crystal per drop is a result of a competitionbetween two processes, nucleation and growth, in a confined volume. Bothprocesses require supersaturation and therefore both nucleation andgrowth are nonequilibrium processes. When the first nucleus forms insidethe drop, it decreases the supersaturation in the surrounding proteinsolution as the crystal grows. If the rate of nucleation is low enough,then the growing crystal will consume enough of the protein in solutionto decrease the supersaturation to the point where another nucleationevent is improbable. Further nucleation is prevented if the time for aprotein to diffuse across a drop is less than the time to nucleate acrystal. Thus combining a small drop volume with the physics ofnucleation and growth, generates negative feedback that acts toautonomously create the ideal dynamical supersaturation profile thatproduces one crystal per drop. Instead of having the negative feedbackimposed externally, as in the Second Harmonic Generation microscopyscheme discussed previously, here the negative feedback is engineeredinto each drop; no external intervention is required. All theengineering goes into identifying the correct combination of diffusiveflux, nucleation rate and drop volume for the emulsions. To completethis screen efficiently, we use polydisperse emulsion droplets asdetailed in the next section.

In this section we calculate the drop volume such that only one crystalis nucleated per drop. Consider a drop that contains a supersaturatedsolution that has not nucleated any crystals. As long as thephysical-chemical environment is constant, the nucleation rate, J [# ofcrystals per unit volume, V, and per unit time, t], will also

be constant and the probability, P, of nucleating a crystal in a drop ofvolume V in an infinitesimal time interval τ is independent of the time,t,

P(t,t+T)=JV _(τ),  (1)

from which it follows that the probability that a drop has not nucleatedany crystals is p(t)=e^(−JVt.) If, by some contrivance, each drop couldonly produce one crystal, then since the probability of notcrystallizing and the probability of crystallizing have to add to one,we have an expression for the average number of crystals per drop as afunction of time;

x(t)=1−e ^(−JVT).  (2)

However, once a drop does nucleate a crystal, the nucleation rate isreduced due to the growing crystal consuming protein in solution andnucleation ceases to be a Poisson process, which makes finding ananalytical solution to the number of crystals per drop as a function oftime a difficult problem.

To address the question of how many crystals nucleate per drop as afunction of drop size, we developed a Monte Carlo simulation in onedimension, a special case for which the drop size and volume are equal.Our approach differs from that taken previously in that our modelexplicitly calculates the spatial-temporal concentration profile withinthe drop. Drops were modeled as a lattice of points, where each pointwas characterized by two quantities; the protein number concentration,c(x,t) [L⁻³], and a binary indicator that signified whether the proteinwas in a crystalline or solution state. The protein was confined in thedrop, meaning that no-flux boundary conditions were imposed on the endsof the lattice. The numerical values used in the model, while within anorder of magnitude of values used in our experiments, were notreflective of any particular protein or physical set of conditions.Rather they were chosen for two purposes. First, to satisfy theassumptions of the theory, i.e. that the rate of crystal growth was muchlarger than the rate of nucleation. Second, to ensure that thesimulations were quick to perform. Thus the diffusion constants andnucleation rates were chosen to be higher than actual values. This meansthat the simulations were faster to perform, but that they conclusionswere not affected as they depend on the ratio of the diffusion rate tonucleation rate and not on their absolute values. Protein concentrationsin solution evolved according to the diffusion equation; δc/δt=D∇²c(t),with D=6×10⁻¹⁰ [m²s⁻¹], the protein diffusion constant. Initially thedrop was homogeneous in protein number concentration, c=1 μm⁻³, at ahigh value of supersaturation, s=83.3, with s=c/c_(s), with c_(s)=0.012,the concentration of the saturated protein solution in equilibrium withthe protein crystal, and with l=1 μm, the size of a lattice site. Ateach time step, there was a finite probability that a randomly chosenlattice site could transform into a crystal with a probability, P, givenby P=Jl³τ and τ=l²/D the simulation time step. In the simulation we usedthe classical nucleation theory expression for nucleation rate,J=sAe^(−B/ln (s)) ² , where A and B are constants such thatP=sR666e^(−350/ln (s)) ² , with R a dimensionless rate coefficient. Theprotein concentration of a lattice site coinciding with the edge of acrystal was increased at a rate proportional to the supersaturationaccording to δ_(c)/δ_(t)=v(c(t)−c_(s))/l, where v=1×10⁻³ [ms⁻¹] is theconstant velocity of crystal growth. Conservation of mass was used atthe boundary between the crystal and solution. The concentration perlattice site in a growing crystal was limited to an arbitrary value ofc_(xtal)=4 to model the effect that protein crystals have a fixeddensity that is of order 100 c_(s). Once a lattice site exceeded thismaximum concentration, the crystal would grow symmetrically, one latticesite to the right and to the left. It was assumed that crystals werestationary once nucleated.

FIGS. 8(A) and 8(B) show the simulation results. Parameters were chosento approximate our experiments; high supersaturation, fast growth, anddrops of order 100 μm diameter. In each case, the simulation begins byinstantly quenching the drop to a supersaturation of 83. The two figuresare taken after nucleation has occurred, but before equilibrium isachieved. In FIG. 8(A), the nucleation rate is low, R=1 in dimensionlessunits. In what follows, time, t, is nondimensionalized by τ=l²/D, anddistance is nondimensionalized by l=1 μm. The red dashed line indicatesthe initial condition, at c=1, while FIG. 8(A) occurs at t=250. Thecrystal that nucleated first is centered at l=19. The width of thecrystal increases as protein from solution is fed into the crystal.Later a second crystal is independently nucleated. The growing crystalsdeplete the protein concentration in the region bordering the crystals.In equilibrium the protein concentration remaining in solution will behomogeneous and equal to the saturation concentration. FIG. 8(B) differsfrom the conditions of FIG. 8(A) in that the dimensionless nucleationrate is higher, R=27. More crystals are formed, even though the durationof the quench at which FIG. 8(B) is recorded, t=50, is less than in FIG.8(A). The protein in solution has obtained the equilibrium value inbetween the two rightmost crystals. A noteworthy observation is thedevelopment of a depletion zone in the neighborhood of each growingcrystal. If the local concentration is reduced sufficiently, then noadditional crystals will nucleate in the depletion zone. The size of thedepletion zone is different between the two figures; therefore thedepletion zone is a function of the nucleation rate J.

FIG. 8(C) shows the average number of crystals per drop as a function oftime obtained from the simulation and compared with a fit to

<x(t)>=x _(∞)(1−e ^(−kt)).  (3)

The simulation conditions were identical to the conditions of FIGS. 8(A)and 8(B); a drop size of 60 μm, and two nucleation rates, R=27 and R=1.The simulation and fit to Equation 3 overlap completely. Equation 2 hastwo fitting parameters, x_(∞), the final number of crystals per drop andk, the non-dimensional rate at which crystals form. FIG. 8(D) shows howthe final number of crystals per drop varies a function of drop size fortwo nucleation rates, R=27 and R=1, while FIG. 8(E) shows how thedimensionless rate, k, varies with size.

FIGS. 8(A) and 8(B) of the concentration profile inside a supersaturateddrop of protein solution during crystallization are suggestive of adepletion zone in the vicinity of a growing crystal in which thesupersaturation is reduced sufficiently such that no new crystals can benucleated. Let w be the width of this depletion zone and let τ be thetime interval for which the average number of crystals nucleated in avolume w^(d), is one, where d is the spatial dimension. Then, fromEquation 1 it follows

Jw ^(d)τ=1,  (4)

which provides one equation relating the depletion zone to thenucleation time. In order for no additional crystals to nucleate in thedepletion zone, the protein in solution must be able to diffuse throughthe depletion zone to the growing crystal, thereby lowering thesupersaturation in the depletion zone, in less than the depletion time.This provides a second equation between the depletion one and nucleationtime,

$\begin{matrix}{\tau = {\frac{w^{2}}{D}.}} & (5)\end{matrix}$

To be self-consistent, we combine Equations 4 and 5, which yields

$\begin{matrix}{w^{d + 2} = {\frac{D}{J}.}} & (6)\end{matrix}$

FIG. 8(D) shows the simulated dependence of the number of crystals perdrop, x_(∞), as a function of drop size in one dimension, d=1, for whichw=(D/J)^(1/3). Examine the curve with the higher nucleation rate, R=27.For small drops, less than drop size ˜9, the number of crystals per dropremains constant at x_(∞)=1. “Small” means V<w^(d), i.e. the time forprotein to diffuse the entire length of the drop is less than thenucleation time, so that after one crystal has been nucleated, itsgrowth causes a negative feedback suppressing further nucleationthroughout the entire drop. As the size of the drop is increased beyondthe depletion zone, w, x_(∞) becomes greater than one and the number ofcrystals per drop grows linearly with drop size. Each nucleation eventproduces a new depletion zone with just one crystal inside. This processrepeats until the entire drop is filled with crystals, each occupying apart of the drop equal to the depletion zone, w. This scenario predictsthat the number of crystals per drop is

$\begin{matrix}{x_{\infty} = {\frac{V}{w^{d}} = {{V\left( \frac{J}{D} \right)}^{\frac{d}{d + 2}}.}}} & (7)\end{matrix}$

The dashed lines in FIG. 8(D) show this behavior; the lines start at theorigin and have slope 1/w. The ratio of nucleation rates in the twoexamples shown in FIG. 8(D) is 27, and as 1/w∝J^(1/3), the prediction isthat the ratio of the slopes of the dashed lines in FIG. 8(D) is 3, asobserved. Furthermore, the width of the depletion zone scales asw∝J^(−1/3). Thus for the drop in FIG. 8(D) with slow nucleation rateR=1, the width of the depletion zone, manifested by the drop size forwhich x_(∞) first becomes greater than one, is predicted to be threetimes greater than the depletion zone of the drop with the fastnucleation rate R=27, as observed in FIG. 8(D).

As the drop volume, V, is increased from zero, the rate, k, ofnucleating one crystal in V will increase linearly with drop volume aspredicted by Equation 1 for Poisson processes

$\begin{matrix}{k_{v} = {\frac{1}{\tau} = {JV}}} & (8)\end{matrix}$

the behavior seen in FIG. 8(E). However, once the drop volume exceedsthe volume of the depletion zone, a crystal will be nucleated somewhereelse in the drop; therefore the frequency at which depletion zones arecreated is

$\begin{matrix}{k = {\frac{1}{\tau} = {{Jw}^{d} = {J^{\frac{2}{d + 2}}{D^{\frac{2}{d + 2}}.}}}}} & (9)\end{matrix}$

Equations 6 and 9 predict that k∝J^(2/3) in one dimension and that kbecomes independent of drop size V, as seen in FIG. 8(E).

The picture that emerges from these simulations and dimensional analysissuggest that nucleation of multiple crystals in a drop is a Poissonprocess. This is an unexpected result as the nucleation rate is notconstant: once the first crystal has nucleated, its growth acts tosuppress further nucleation. However, we argue that each nucleationevent creates a depletion zone in which it is only possible for onecrystal to exist. Therefore, each nucleation event is an independentprocess. In effect, each drop can be thought of as being partitionedinto x_(∞) smaller, independent drops of volume w^(d)=V/x_(∞) thatnucleate with rate k (FIG. 8 F). This justifies Equation 3, and explainswhy in FIG. 8(C), the number of crystals per drop as a function of timeis an exponential, a result of a Poisson process.

The degree to which growing crystals create depletion zones is expectedto be greatest in one dimension. For example, in one dimension noprotein can be replenished in the gap between two crystals, while inhigher dimensions, protein will diffuse into the gap between twocrystals along the directions perpendicular to the line connecting thecenter of the crystals. Nevertheless, we expect the same general trendsobserved in 1D to carry over to 2D and 3D. In particular, in dimension dwe expect there will be a drop volume

${V_{d} \sim \left( {D\text{/}J} \right)^{\frac{d}{d + 2}}},$

below which only one crystal will be nucleated per drop.

3. Crystal Emulsions

To yield identical crystals in sufficient quality and quantity forserial crystallography, we use a two step method. We first identify theappropriate drop volume to consistently nucleate one crystal per drop.For this we intentionally created emulsions in a batch process thatyielded a polydisperse size distribution, ranging from a few microns toa few hundreds of microns in diameter (FIG. 9A-C). Such a polydisperseemulsion allowed us to identify the critical diameter in a singlescreening experiment. We then used microfluidics (FIG. 9D) to producemonodisperse emulsion droplets (FIGS. 9E, 9F), which we used to growidentical crystals in the serial X-ray diffraction chip, as described insection 5 below. For the purposes of this example, however, the fullexperimental sequence will only be reported for glucose isomerase, i.e.,whereby crystals were grown in the serial diffraction chip, X-ray datawere acquired, and the structure was solved.

All crystals were grown in emulsion droplets stabilized againstcoalescence with a 2% v/v solution of PFPE-PEG-PFPE non-ionic triblocksurfactant “E2K0660” in Novec™ HFE-7500 fluorinated oil (from 3M). ThePFPE-PEG-PFPE surfactant was synthesized as previously described inHoltze et al., (2008) Lab on a Chip, 8(10), 1632-9. PFPE is aperfluorinated polyether, —CF(CF₃)CF₂O—, and polyethyleneglycol (PEG) is—CH₂CH₂O—. Note that a commercial surfactant is now available from RANBiotechnologies, Beverly, Mass., USA. Novec™ HFE-7500 is2-(Trifluoromethyl)-3-ethoxydodecafluorohexane. We chose a fluorinatedoil and a fluorinated surfactant to minimize interactions withbiological molecules. Fluorocarbon and hydrocarbon oils do not mix witheach other, nor do they mix with water. In particular, the PFPE-PEG-PFPEsurfactant in HFE-7500 oil system has been shown to have excellentbio-compatibility. To confirm that it is compatible with proteincrystallization, we tested it with five crystallization model proteins(FIG. 9 and Table 1). All five model proteins have previously beencrystallized by vapor diffusion and a structure derived from X-raycrystallography deposited in the PDB.

TABLE 1 formula net charge weight Isoelectric point (pI) in crystalLysozyme 14.3 kDa 11.3 (from (Wetter & positive Deutsch, 1951)) Trypsin 24 kDa 10.1-10.5 (From (Walsh, positive 1970)) Concanavalin 76.5 kDa4.5-5.5 (multiple isoforms, negative A (3mer) see (Entlicher et al.,1971)) Glucose  173 kDa 3.95 (from (Vuolanto negative isomerase (4mer)et al., 2003)) D1D2 26.8 kDa 10.6 (theoretical pI, from positive(heterodimer) (ProtParam tool, 2013))

To adopt a published vapor diffusion recipe into our emulsion format, wehad to perform a set of pre-experiments. In traditional vapor diffusion,a small volume of protein solution is mixed with the same amount ofprecipitant and then sealed into a container together with a largereservoir of precipitant. The diluted protein-precipitant dropequilibrates through vapor phase diffusion with the reservoir, resultingin a concentration increase of all components in the drop byapproximately a factor of two. All previously published crystallizationrecipes had been optimized to nucleate only a few crystals permicroliter. Our emulsion droplets have volumes of a few picoliters tonanoliter each. As the probability of nucleating a crystal isproportional to the sample volume, we had to increase nucleation ratesby at least two orders of magnitude. We thus prepared vapor phase andmicrobatch crystallization trials around the literature recipes andoptimized the vapor recipes toward nucleating crystal showers ofappropriate density. When attempting to crystallize a novel proteintarget through screening crystallization conditions such crystal showersare usually considered a first hit and the conditions are later refinedextensively to grow the largest possible crystal. When using the methodpresented here on a novel protein target, the polydisperse emulsionscreen would directly follow onto the initial hit finding, and thereforeeliminate the reverse engineering step of converting an optimized vaporphase recipe back to a recipe that grows crystal showers.

Polydisperse emulsions were then prepared by mixing 2 μL proteinsolution with 2 μL precipitant in a 150 μL PCR test-tube. Immediatelyafter mixing, we added 30 μL 2% v/v solution of PFPE-PEG-PFPE surfactant(E2K0660) in HFE-7500 fluorinated oil. Polydisperse emulsions wereformed by gently agitating the vial by hand until droplets became toosmall to be resolved by eye. This procedure typically gave dropletsranging from a few microns to a few hundreds of microns in diameter(FIG. 9A-C). Aqueous droplets were less dense then the immersingfluorinated oil, so droplets rose (“creamed”) to the top of the vialwithin a minute. The emulsion cream was then loaded into rectangularglass capillaries (VitroTubes from VitroCom, Mountain Lakes, N.J., USA)and sealed with 5 minute epoxy to prevent evaporation. Crystallizationwas monitored over the course of a week. All compounds and proteins fromcommercial sources were used as received without further purification.The molecular weight and the net charge of the proteins duringcrystallization, as derived by the isoelectric point, are summarized inTable 1.

Lysozyme was crystallized by encapsulating 30 mg/ml Lysozyme, 100 mMsodium acetate, pH 4.8, 12.5 wt % PEG 8000, 5 wt % NaCl finalconcentration into droplets and then incubating them at 6° C. for 36hours until all droplets had nucleated crystals. This recipe was derivedfrom a vapor phase recipe mixing 20 mg/ml lysozyme in 100 mM sodiumacetate pH 4.8 with an equal volume 10% (w/v) NaCl, 100 mM sodiumacetate pH 4.8, and 25% (v/v) ethylene glycol.

Glucose isomerase crystals were grown at room temperature (˜25° C.) bypreparing a crystallization batch with final concentrations of 30 mg/mlglucose isomerase from Streptomyces rubiginosus (from Hampton Research),100 mM ammonium sulfate, pH 7.0, 20 wt % PEG 10,000 in a 1:1 ratio (allfrom Sigma Aldrich). The initial vapor phase crystallization conditionwas taken from the Hampton Research data sheet as mixing 20-30 mg/mlglucose isomerase with 10 to 15% (w/v) PEG 4000-8000, 200 mM salt, pH6.0-9.0.

Trypsin was crystallized by mixing 60 mg/ml trypsin (Sigma T-8253) frombovine pancreas in 10 mg/ml benzamidine, 3 mM CaCl₂, 0.02 wt % sodiumazide with 100 mM NaPO₄, pH 5.9, 5.1 M ammonium acetate (all SigmaAldrich). In this system we observed crystals in the range of pH 5.9 topH 8.6, with higher pH values having much higher nucleation rates. Thisrecipe was derived from a vapor phase recipe mixing 60 mg/ml trypsin in10 mg/ml benzamidine, 3 mM calcium chloride, and 0.02% (w/v) sodiumazide with an equal volume of 4% (w/v) PEG 4000, 200 mM lithiumsulphate, 100 mM MES pH 6.5, and 15% ethylene glycol.

Concanavalin A was crystallized by mixing 25 mg/ml concanavalin A typeIV from Canavalia ensiformis in 10 mM tris hydrochloride, pH 7.4 with100 mM tris hydrochloride, pH 8.5, 8 wt % PEG 8,000 in a 1:1 ratio (allfrom Sigma Aldrich). For this we first set-up vapor phase and microbatchtrials of 20 mg/ml Concanavalin A in 10 mM TRIS pH 7.4 against the 50conditions in the Hampton Crystal Screening Kit. From this screen wechoose condition 36, with 100 mM tris hydrochloride, pH 8.5, 8 (w/v) PEG8000, as this condition grew crystals in both vapor phase and microbatchtrials.

D1D2, the sub-complex from the human snRNP spliceosome core particle(PDB entry 1B34), crystallized over 72 hours at room temperature bypreparing a crystallization batch with final concentrations of 6 mg/mlD1D2, 62 mM sodium citrate pH 5.2, 125 mM ammonium acetate, 9 vol %glycerol, 26 wt/vol PEG 4,000 (all Sigma Aldrich). D1D2 was purified aspreviously reported. D1D2 was first crystallized by Kambach et al. invapor phase by mixing equal volumes of 6 mg/ml D1D2 in 20 mM sodiumHEPES pH 7.5, 200 mM sodium chloride and 6 mM dithiothreitol and 100 mMsodium citrate pH 5.6, 200 mM ammonium acetate, 15% glycerol, 25% PEG4000.

All globular proteins, concanavalin, glucose isomerase and trypsin,crystallized readily in vapor diffusion, microbatch, and the emulsionsystem. The heterodimer D1D2 formed crystals in vapor phase and theemulsion system only. In microbatch, a thick protein skin grew at thedroplet interphase potentially depleting all the protein from the drop.We thus conclude that the PFPE-PEG-PFPE surfactant system is well suitedto protect protein from absorbing to the fluorooil-water interface andto stabilize emulsions, making it ideal for crystallization trials.

All initial crystallization experiments were performed at roomtemperature. However, a particular protein may become unstable at toohigh or too low temperatures. Also, many proteins like lysozyme havetemperature sensitive nucleation rates, which one might like to exploitin crystallization trials. An ideal surfactant oil system can hence beused in a large temperature range. To test for temperaturecompatibility, we prepared crystal emulsions from the PFPE-PEG-PFPEsurfactant in HFE-7500 oil, sealed them into rectangular glasscapillaries and incubated them in a water bath at 4° C. and at 40° C. Wefound the emulsion droplets to be stable for at least two weeks at thosetwo temperatures.

Finally, to yield identical crystals in sufficient quantity for serialcrystallography, we employed microfluidics to produce monodisperseemulsion droplets. For this, we simply selected the dropmaking chipappropriate to make drops of the desired diameter and used thecrystallization recipe from the preceding polydisperse emulsion screenwithout further modification. We produced drops in a co-flow geometrydesigned such that the protein solution and buffer do not mix in thelaminar flow upstream of the dropmaker (FIG. 9D). Typically, injectionof the oil-surfactant mixture proceeded at 600 μL/hr, while both proteinand precipitant streams were pumped at equal flow rates of 300μ/hr toco-encapsulate both in a 1:1 mixture. Upon droplet formation, mixinginside each droplet proceeds within less then a second due torecirculating flow that arises from shearing interactions of the fluidinside the drops with the stationary wall. These monodisperse emulsiondroplets were then injected into and incubated in the diffraction chipto grow crystals for the X-ray diffraction experiments.

To monitor crystallization, we stored emulsion droplets in two differentsystems. Firstly, polydisperse emulsions were usually sealed intorectangular glass capillaries, which prevented water and oilevaporation. Secondly, as our diffraction chip was made from a polymermaterial, we exploited its permeability to water vapor by slowly lettingdroplets shrink by permeation of water from the drops into the oil andalso from the drops through the thin, polymer-based chip. Waterpermeation across the polymer foil decreases linearly with increasingfoil thickness and decreased permeability of the material. In case ofthe 50 to 75 μm thick cyclic-olefin-copolymer (COC) sheets used here,the evaporative water loss amounted to a few percent per hour. Whenwater evaporates from the drop, the solute concentrations inside thedrop increase and hence the protein supersaturation also increases. Asthis corresponds to an increased nucleation rate, one would expect toyield a larger fraction of droplets with multiple crystals. We did notobserve such an effect and attribute this to the fact that once thefirst crystal nucleates, its subsequent growth reduces thesupersaturation of the solution enough to prevent another crystal fromnucleating. We consistently achieved one crystal per drop, which arguesfor the robustness of the method. Once all droplets had nucleatedcrystals, we immersed the capillary/chip into an oil bath to preventfurther evaporation. Alternatively we achieved equally good results withstoring chips in a water bath to which a vial containing an oilreservoir was connected to the chip and all other inlets where sealed.

4. X-Ray Semi-Transparent Chip Fabrication

Looking at FIGS. 10 and 10D, chips 50 were sealed, which is colloquiallyreferred to in the thermoplastic industry as ‘lidding’, by bondingcyclic-olefin-copolymer (COC)-foil or Kapton® polyimide-foil 51, 53 toboth sides of the thin poly(dimethylsiloxane) (PDMS) slab 52 containingthe channels 59 (FIG. 10A3). (The channels could also be as shown inFIGS. 3A to 3C.) PDMS (Sylgard® 184 from Dow Corning) with ratio 1:5 ofcuring agent to base was molded on a standard SU8-master 55 by squeezingthe uncured PDMS resin into a thin film using a glass plate 57 and aweight 58. To facilitate release of the PDMS film, the master wassurface treated with a fluorophilic coating by spin-coating 1:20 CytopCTL-809M in CTsolv.100E (both Bellex International) onto the master. Wethen baked the wafer for 1 hour at 150° C. We placed a 30 μm thick Mylarfoil 56 (DuPont) between PDMS 52 and glass 57 to allow for easy removalof the glass slide after PDMS curing. We pre-cured the PDMS for 4 hoursat room temperature before we removed the weight and transferred thecomplete stack into the oven to drive the curing reaction to completionat 72° C. for another hour.

We either used COC (TOPAS® 5013 cyclic olefin linear olefin copolymerfrom Advanced Polymers) or Kapton® poly(4,4-oxydiphenylenepyromellitimide) (American Durafilm), depending on experimentalrequirements. COO is more brittle than Kapton®, but has a lower watervapor permeability. The thinnest commercial COO we used was 25 μm thinTOPAS®, while Kapton® as thin as 8 μm can be purchased as bulk foil. Wechemically bonded either foil substrate 51, 53 to the featured PDMS 52using a silane coupling chemistry. In brief, both foil and PDMS areactivated in an oxygen plasma and then incubated for 25 minutes in anaqueous solution of a different silane each; 1 vol % of3-aminopropyltrimethoxysilane (APTMS, 97% from Aldrich), and 1 vol % of3-glycidoxypropyltrimethoxysilane (GPTMS, 98%, from Aldrich). The twosilanes are such that they can form an epoxy bond when brought incontact. Upon removing foil and PDMS from the batch, we dried both witha stream of nitrogen gas and then carefully brought them in contactusing tweezers to prevent trapping air bubbles between both layers. Thechip 50 was then incubated in the oven at 72° C. for 1 hour to maximizechemical cross-linking. The process was repeated to lid the other sideof the chip. Upon assembly, the chip 50 was surface treated with afluorophilic coating to prevent protein interaction with the channelsurface. For this, 1:20 Cytop CTX-109AE in CTsolv 100E (both BellexInternational) was dead-end filled into the chip by plugging all outletsand slowly injecting the Cytop solution through the inlet into the chip.This causes gas bubbles trapped inside the chip to become pressurizedwhich promotes the gas to dissolve into solution and also to permeateacross the chip walls to result in a completely filled, bubble freedevice. The chip was then incubated at 90° C. for at least 12 hours toevaporate the solvent away and also to accelerate chemical cross-linkingbetween fluoropolymer and chip surface.

In the non-limiting example of FIG. 10A(3), the foil cover 53 has abottom to top thickness of about 25 μm, the PDMS layer 52 has a bottomto top thickness of about 80 μm with channels of a bottom to top heightof about 75 μm, and the foil cover 51 with holes has a bottom to topthickness of about 50 μm. However, a horizontally oriented single layerchip 50 a is also possible as shown in the non-limiting example of FIG.10D in which the single layer of foil 51 a has a bottom to top thicknessT of about 25 μm. The PDMS storage section 52 a with channels 59 a isoriented vertically in the layer of foil 51 a as shown in FIG. 10D. Aplurality of storage sections can be oriented vertically in the layer offoil. The embodiment of FIG. 10D could provide for improved efficiencyby reducing the time to optimize conditions for the largest crystal.

5. In-Situ Diffraction

We mounted the X-ray transparent chip into a custom acrylic frame tocollect diffraction data (FIG. 11). The acrylic frame was cut to shapefrom 3 mm thick acrylic sheet using a 40 W CO₂ Hobby Laser cutter with a1.5″ focus lens (Full Spectrum Laser). To create ports into thefoil-chip we drilled through holes into the acrylic frame with the lasercutter. Blunt needle pins (23 gauge) were then placed into the holes andglued into position with 5 minute epoxy. We connected #30 AWGpoly(tetrafluoroethylene) (PTFE) tubing (Cole Palmer) to the pins usingPDMS cubes with through holes punched into them using 0.75 mm HarrisonUni-Core biopsy punches (Ted Pella). Buna O-rings, 70 durometer, size002 (McMaster Carr) were then used to seal the foil-chip to the hollowmetal pins. For easy alignment the o-rings were fit into a 1 mm thickpoly(ethyleneterephthalate) (PET) spacer that also was fabricated withthe laser cutter. X-ray semi-transparent foil-chips were mounted into aframe for the duration of each experiment. Each frame was held togetherby 10 self-tapping 3/16″ Pan Head 2-28 Phillips screws (McMaster Carr)to lock the chip into position and to minimize flow induced inside thechip from mechanically bending the thin foil chip. To mount theframe-chip assembly in the synchrotron we machined a stainless steeladapter that a frame could be mounted onto using two screws (FIG. 11B).

For the proof of principle experiment, we fabricated an X-raysemi-transparent chip with the “dropspot” geometry that can hold up to8000 emulsion droplets in cavities with 150 μm diameter each (FIG. 10B&C). The fluorinated oil has a density of 2 g/mL, while the water dropshave a density of 1 g/mL. Thus there is a strong tendency for the dropsto float to the top of the oil, or “cream”. Surface tension forcesarrest droplets in a cavity and prevent them from creaming to one sideof the chip. We produced a monodisperse ˜110 μm diameter emulsion of 30mg/ml glucose isomerase, 100 mM Ammonium Sulfate, pH 7.3, 20 wt % PEG 10k MW final concentration in a standard dropmaker (FIG. 10D). Dropletsexiting the dropmaker were immediately routed into the X-raysemi-transparent serial crystallography chip by simply plumbing thedropmaker outlet into the dropspot inlet (FIG. 11A). After the dropspotchip was loaded, we dead-end plugged its outlet except for one inletwhere we kept HFE-7500 oil entering the chip using hydrostatic pressureto compensate for oil evaporation from the chip. We incubated the chipat room temperature for three days and monitored crystallization, beforetransferring into a water bath to prevent further evaporation. By then,most droplets had shrunken to about ˜90 μm diameter and more than 90% ofthem had nucleated a single crystal. Crystals grew to about 50 μm by 40μm by 30 μm in size at room temperature (˜25° C.).

X-ray diffraction data were collected at Cornell High Energy SynchrotronSource (CHESS), beamline F1 (λ=0.9179 Å, E=13.508 keV), using a 100 μmmonochromatic X-ray beam from a 24-pole wiggler. The chips were mountedat a distance of 200 mm from an Area Detector Systems Corporation (ADSC)Quantum 270 (Q270) detector, corresponding to a largest inscribed circleof resolution of 1.4 Å. The detector face was oriented perpendicular tothe beam. For selected crystals within the chip, data sets werecollected at room temperature (˜25° C.). Each recorded data setcomprised 10 frames, for a total of 10° oscillation. Each imageconsisted of a 5 second exposure with a 1° oscillation step size. Atotal of 1520 images were collected from 152 glucose isomerase crystalsin three different dropspot chips.

6. X-Ray Structure Determination

The software HKL-2000 was used to index, refine, integrate and scaleeach 10° data set before merging. Parameters including unit-cell size,chi-squared values, resolution, mosaicity, and completeness wereevaluated for every partial data set during the indexing and scalingprocess. From these partial data sets, with 1520 frames total, weselected 262 frames from 72 crystals by rejecting frames with a mosaicspread higher than 0.1° and chi-squared x and y (corresponding todiscrepancy between observed and predicted spot positions) above 2. Someframes were later rejected because of poor scaling statistics; the finaldata set included 248 frames.

Glucose isomerase crystals were determined to have a space group ofI222, and diffracted to an average of 2 Å; an example image is shown inFIG. 11E. In some crystals, diffraction extended to 1.4 Å, with a mosaicspread of 0.04.

The 248 selected frames were scaled together using Scalepack (HKLResearch) and merged with Aimless. The limiting resolution of 2.09 Å waschosen as that at which CC1/2 dropped below 0.5. Statistics are given inTable 2. The merged data set covered 93% of reciprocal space, suggestingthat preferred orientation of the crystals was not a major problem. Theglucose isomerase structure was readily solved by molecular replacementwith Molrep using the structure previously determined at 1.90 Åresolution (PDB ID: 8XIA), with waters removed. Prior to refinement, werandomly flagged 5% of the reflections for R_(free) analysis.

Structure refinement was carried out through multiple iterations ofRefmac, refining atomic coordinates and isotropic B-factors. 2Fo-Fc andFo-Fc electron density maps were generated after each refinement step,and further refinement was carried out by manual inspection using Coot.In the refinement process, two disordered N-terminal residues wereremoved, as well as a bound sugar molecule present in the model but notin the crystal, and 124 water molecules were added. Final refinementgave R and R_(free) of 14.4% and 17.5%, respectively. Completeprocessing statistics are given in Table 3. FIG. 12 shows the quality ofthe final refined structure.

TABLE 2 Processing results of merging the 248 frames obtained from 72glucose isomerase crystals. Values in parentheses refer to the highestresolution bin (2.15-2.09 Å). Precipitant composition 100 mM ammoniumsulfate pH 7.0 + 20 wt % PEG 10,000 Space Group I222 Unit-cellparameters (Å) a = 93.94 b = 99.47, c = 102.85 Resolution range (Å)49.7-2.09 (2.15-2.09) No. of unique reflections 26699 (2075) Redundancy8.2 (8.1) Completeness (%) 93.2 (94) R_(merge) (%) 0.191 (0.686) <I/σ(I)> 7.8 (4.1) Mosaicity (°) 0.03-0.1

TABLE 3 Refinement and model statistics for glucose isomerase. Values inparentheses refer to the highest resolution bin. Resolution range (Å)49.7-2.09 (2.14-2.09) Reflections used: working, total 25395, 26685(1879, 1974) Completeness (%) 92.4 (93.6) R(working)/R_(free)0.144/0.174 (0.186/0.227) RMSD, bond lengths (Å) 0.019 RMSD, bond angles(°) 1.93  No. of protein/other atoms 3034/126 (non-hydrogen) Mean Bvalue, all atoms (Å²) 17.6   Ramachandran statistics (%); 97.13, 2.35,0.52 favored, allowed, outliers R and R_(free) are calculated using Σ|Fo| − |Fc|/Σ |Fo| for the working and free-set reflections,respectively

7. Conclusion

In this example, we present a technology that optimizes the kinetics ofcrystallization, eliminates crystal handling, eliminates cryoprotectionand simplifies collection of diffraction data for structural biology. Inthis example, we developed processing methods for proteincrystallization that follow the ideal kinetic pathway of slowlyincreasing supersaturation until a single crystal nucleates and thenreducing supersaturation so that one crystal grows slowly to allowannealing of defects. Sample volume is not a thermodynamic variable inphase equilibrium, but since crystallization is a non-equilibriumprocess, volume plays a key role in determining the kinetics ofcrystallization. We believe using a combination of simulation, theoryand experiment that selecting the appropriate droplet diameter, w,guarantees that only one crystal per drop will form when the drop volumeV<˜(D/J)^(d/2+d.) We identify the critical drop diameter for aparticular crystallization condition in a single experiment by using apolydisperse emulsion with droplets ranging from a few micrometers to afew hundreds of micrometers in size. These polydisperse emulsions can bemade with ease within seconds using only a pipette and a test tube. Theprobability of crystallization is proportional to drop volume. As we usedrops of order 1 nanoliter, which are smaller drops than employed byother methods, the nucleation rates and supersaturation that we use arehigher than usual.

Employing these kinetic processing methods, we grew monodispersecrystals compartmentalized in emulsion droplets, with one crystal perdrop. Monodisperse, microfluidically produced drops of supersaturatedprotein solutions were stored on chip and slowly concentrated as waterpermeated through the thin foil chip. Single crystals per drop werenucleated and grown on-chip in identical conditions. While cyro-cooledcrystals can be stored almost indefinitely, the crystals grown andstored in our chips are stable for several weeks when the chips arestored in a water bath connected to an oil reservoir, which preventsevaporation and hence drying out. The chip for nucleating crystals wasthin enough to be X-ray semi-transparent and diffraction patterns werecollected from these crystals on-chip at room temperature. The structureof glucose isomerase was solved and refined at 2.09 Å resolution, to anR_(cryst)/R_(free) of 0.144/0.174, using merged diffraction datasetsfrom 72 crystals of about 50 μm by 40 μm by 30 μm in size.

Diffraction from room temperature crystals stored on the chip in whichthey were nucleated and grown has many advantages over traditionaloff-chip cyroprotected crystals. On-chip diffraction means the crystalsare not removed from their mother liquor, which can lead to dehydrationand osmotic shock of the crystals and the generation of stress andstrain. Room temperature diffraction eliminates the laborious step ofcryoprotection and has the additional effect of lowering the mosaicityas cryoprotection generates stresses due to changing solvent conditionsand temperature induced volume changes. Our chip can be inexpensivelymass produced and is simple to operate without the need of controllingvalves. We envision a chip that uses temperature and concentrationgradients to discover optimal crystal growth conditions wherein crystalswould be grown at the optimal conditions to create a stream of tinycrystals that would be serially conveyed to a part of the chip withultra thin windows for in-situ diffraction.

Example 4

In this example, we illustrate fabrication of an X-ray transparentcrystallization device.

Harvesting crystals from microfluidic devices damages crystals, becausestresses introduced by environmental changes and mechanical manipulationcan strain or destroy protein crystals. We seek to overcome thislimitation by leaving the crystals on the chip, bathed In the motherliquor from which the crystals were produced. We do this by making thechips so thin that x-rays pass through the chip without significantscatter. We use tooling to produce thin foil microfluidic devices. Thisenables us to produce thinner chips.

A microfluidic device of this example greatly improves the determinationof protein structure, which is necessary for fundamental knowledge,medical research and pharmaceutics. The chip produces crystals usingnovel crystallization optimization protocols. Reducing the chip in sizeto 150 microns will render it x-ray transparent, permitting structuresto be obtained from crystals on chip. The non-limiting example crystalchip has two integrated layers, one containing protein and the otherbuffer solutions, linked by a selective membrane. Current technologiesrequires crystals to be harvested manually, damaging the fragilecrystals. To manufacture the thin crystal chip, we use a thermo-pressfor embossing the microfluidics into biocompatible plastic films.

The microfabrication tooling includes a laminator and a thermal pressthat operates in the 1 to 10 psi pressure ranges. The custom toolingincludes a pressure sensor, piston system, temperature regulator (up to300° C.), and a vacuum pump. The tool may be configurable to work in twodifferent ways. First, actuated by pressurized air, two heated metalplates can be pressed against each other to produce thin plastic plaquesand foils. Second, a vented chamber can be sandwiched between the heatedplates to pull and press a thin foil onto an embossing master using thevacuum.

An example sequence of steps for manufacturing an X-ray transparentcrystallization device is as follows. Step 1 comprises manufacturing amaster, i.e. by micromilling a master form. Step 2 comprises casting anegative mold insert with PDMS on the micromilled master form. Step 3comprises assembling the negative mold insert and a polymer (e.g., COC)foil into an aluminum tool holder prior to evacuation. Step 4 comprisestightly sealing the tool holder and heating above a glass transitiontemperature of the polymer foil. Step 5 comprises the application ofpressurized nitrogen into the interior of the tool holder. Step 6comprises demolding the polymer foil after cooling.

A first negative mold insert can be created for embossing a microfluidicchannel having a plurality of microwells in a first polymer foil. Asecond negative mold insert can be created for embossing one or morereservoirs in a second polymer foil. A fastening system can be used forattaching the first polymer foil to a first side of a dialysis membraneand for attaching the second polymer foil to a second opposite side ofthe dialysis membrane creating a structure as in FIG. 10(A) 3.

Example 5

In this example, we illustrate X-ray transparent differential permeationchips. FIG. 13 shows cross sectional views of an example single membranedifferential permeation X-ray chip 60 and an example double membranedifferential permeation X-ray chip 70 according to the invention. Theconcept is to make the membrane have a wedge-shape cross-section togenerate a gradient of permeation flux. The non-limiting examplemembranes in FIG. 13 use a permeation membrane attached to the X-raychip in a reversible way so that the differential permeation membranecan be removed before shooting crystals. The devices of FIG. 13 operateby dipping the assembly into different reservoir solutions, i.e., firstinto a high salt buffer, or exposed to air to shrink drops and then intowater to swell drops. The double membrane differential permeation X-raychip has the permeation control on both sides for a two times transferrate. The differential permeation membrane could use an elastomeric(e.g., PDMS) membrane, to only permeate water and non-polar solutes.

In the single membrane differential permeation X-ray chip 60 of FIG. 13,cyclic-olefin-copolymer (COC)-foil or Kapton® polyimide-foils 61, 63 arebonded to both sides of a poly(dimethylsiloxane) (PDMS) layer 62containing the channels 69. Fluid ports 67 are in fluid communicationwith the channels 69. A permeation barrier 64 is bonded to the foil 62.A differential permeation membrane 65 of varying thickness creating awedge-shape cross-section is bonded to the foil 61. The membrane 65 maycomprise a dialysis membrane, or the membrane 65 may comprise a membranepermeable to water, or the membrane 65 may comprise a polyethersulfone,or the membrane 65 may comprise regenerated cellulose or celluloseester. The membrane 65 may be hydrophilic. The foils 61, 63 may have athickness of about 5 μm to about 100 μm, the PDMS layer 62 may have abottom to top thickness of about 50 μm to about 150 μm with channels ofa bottom to top height of about 45 μm to about 145 μm, and the membrane65 may have a thickness varying from about 5 μm to about 100 μm.

In the double membrane differential permeation X-ray chip 70 of FIG. 13,cyclic-olefin-copolymer (COC)-foil or Kapton® polyimide-foils 71, 73 arebonded to both sides of a poly(dimethylsiloxane) (PDMS) layer 72containing the channels 79. Fluid ports 77 are in fluid communicationwith the channels 79. Differential permeation membranes 75, 76 ofvarying thickness creating a wedge-shape cross-section are bonded to thefoils 71,73. The membranes 75,76 may comprise a dialysis membrane, orthe membranes 75,76 may comprise a membrane permeable to water, or themembranes 75,76 may comprise a polyethersulfone, or the membranes 75,76may comprise regenerated cellulose or cellulose ester. The membranes75,76 may be hydrophilic. The foils 71, 73 may have a thickness of about5 μm to about 100 μm, the PDMS layer 72 may have a bottom to topthickness of about 50 μm to about 150 μm with channels of a bottom totop height of about 45 μm to about 145 μm, and the membranes 75,76 mayhave a thickness varying from about 5 μm to about 100 μm.

Example 6

In this example, we illustrate an X-ray transparent chip storagecontainer. Referring to FIG. 14, there is shown a cross sectional viewof an X-ray chip storage container 80 according to the invention. Thestorage container prevents both: uncontrolled loss of oil and buffer.FIG. 14 shows an all in one vial 81, but it can be separate vials aswell. Using thin X-ray transparent foils comes at the expense of thechip being sensitive to drying out over a few hours and hence long-termstorage is a concern. We use a special vial 81 as shown in FIG. 14 to beable to transport (such as by mail) loaded chips.

The components include a vial 82 of oil connected to the X-ray chip 83,with all other chip fluid outlets being blocked. The vial 82 has anopening 91 such that hydrostatic pressure can push fresh oil into thechip to replace oil that has evaporated. The opening 91 can be sealedduring shipment and opened (such as by breaking a frangible seal) beforeuse. The chip 83 has cyclic-olefin-copolymer (COC)-foil or Kapton®polyimide-foils 84 bonded to both sides of a poly(dimethylsiloxane)(PDMS) layer 85 containing channels 86. The chip 83 itself is placed inan aqueous bath 88 of buffer to prevent water evaporation. Fluid conduit89 is in fluid communication with a reservoir 94 of the layer 85 and thevial 82 of oil to allow transfer of the oil by way of hydrostaticpressure. Taking these measures extends the shelf life to at least amonth for an X-ray chip.

Example 7

In this example, we illustrate an apparatus for controlling a reactionor a phase transition in which hydrostatic pressure driven flow is usedto control dialysis. Non-limiting examples of the reaction are one ormore of the following: steady-state and self-assembly reactions at orfar from equilibrium; perturbation analysis of reaction networks; cellsynchronization; cell and tissue differentiation; and/or chemostatreactions with cells and cell populations. Non-limiting examples of thephase transition are one or more of the following: crystallization andco-crystallization of small molecules, biological macromolecules,colloids and combinations thereof; liquid crystal phase transitions;gelation; liquid-liquid separation; protein folding; and/or DNA meltingor condensation.

Looking at FIG. 15, the non-limiting example apparatus 110 forcontrolling a reaction or a phase transition includes a microfluidicdevice 112 such as that shown in FIG. 1. In FIG. 15, only the storagelayer 114 and the reservoir layer 116 of the microfluidic device 112 areshown for ease of illustration. The microfluidic device 112 of theapparatus 110 includes a reservoir layer defining a reservoir; adialysis membrane disposed on the reservoir layer; a wetting controllayer disposed on the membrane; and a storage layer disposed on thewetting control layer. The wetting control layer and the storage layerdefine a microfluidic channel comprising an upstream portion, adownstream portion, a first fluid path in fluid communication with theupstream portion and the downstream portion, and a storage wellpositioned within the first fluid path. The wetting control layerincludes a fluid passageway in fluid communication with the storage welland the membrane. The wetting control layer is capable of wetting afirst fluid introduced into the microfluidic channel, the first fluidcomprising a hydrophilic, lipophilic, fluorophilic or gas phase as thecontinuous phase in the microfluidic channel.

The apparatus 110 also includes a source of oil, vial 122, in fluidcommunication via a conduit 123 with the microfluidic channel of themicrofluidic device 112; and a source of an aqueous fluid, vial 126, influid communication via a conduit 127 with the reservoir of themicrofluidic device 112. The vials 122, 126 containing the oil andbuffer have open tops such that hydrostatic pressure regulates transportfluxes across the membrane of the microfluidic device 112. Notepressures p_(storage) and p_(reservoir) in FIG. 15. Alternatively, apressure controller can regulate transport fluxes across the membrane.Storage outlets 133 of the microfluidic channel can be blocked, and anoutlet vessel 135 of the reservoir can be open. A microscope 141 is usedfor monitoring the reaction or the phase transition in the microfluidicdevice 112.

Example 8

Growth kinetics, the relationship between cell growth rate and nutrientsupply, plays a vital role in the understanding of cell function. Achemostat is one example device for the study of the growth kinetics ofmicroorganisms. Chemostats can maintain a constant population of amicroorganism in a state of active growth. This may be done byperiodically substituting a fraction of a culture with an equal volumeof fresh, sterile, chemically defined growth medium. However, one majordifficulty with a chemostat is the need to continuously supply medium ascontinuous cultures can run for days or weeks at steady state.

A microfluidic chemostat of the present invention offers a way toaddress the difficulties relating to conventional continuous culturesystems. A microfluidic chemostat of the present invention can run forlong periods of time consuming much less media.

We prepared a dialysis chip as in Example 1 with the microfluidicchannels 40 of FIG. 3. This dialysis chip can be used for studying thebiome. We used this dialysis chip with microfluidic channels 40 as achemostat that kept alive yeast populations for a week. The deviceoperated at constant conditions—continuously supplying nutrients andremoving waste and yeast, thereby maintaining a constant population.FIG. 16 shows yeast populations in wells having a volume of 20nanoliters each at a time lapse of approximately one week.

Thus, the invention provides microfluidic devices for investigatingcrystallization, and microfluidic devices for controlling a reaction ora phase transition.

Although the invention has been described in considerable detail withreference to certain embodiments, one skilled in the art will appreciatethat the present invention can be practiced by other than the describedembodiments, which have been presented for purposes of illustration andnot of limitation. Therefore, the scope of the appended claims shouldnot be limited to the description of the embodiments contained herein.

1. A microfluidic device comprising: a reservoir layer defining areservoir; a membrane disposed on the reservoir layer; a wetting controllayer disposed on the membrane; and a storage layer disposed on thewetting control layer, wherein the wetting control layer and the storagelayer define a microfluidic channel comprising an upstream portion, adownstream portion, a first fluid path in fluid communication with theupstream portion and the downstream portion, and a storage wellpositioned within the first fluid path, wherein the wetting controllayer includes a fluid passageway in fluid communication with thestorage well and the membrane, and wherein the wetting control layer iscapable of wetting a first fluid introduced into the microfluidicchannel, the first fluid comprising a hydrophilic, lipophilic,fluorophilic or gas phase as the continuous phase in the microfluidicchannel.
 2. The microfluidic device of claim 1 wherein: the membranecomprises a dialysis membrane.
 3. The microfluidic device of claim 1wherein: the membrane comprises a membrane permeable to water.
 4. Themicrofluidic device of claim 1 wherein: the membrane comprises apolyethersulfone.
 5. The microfluidic device of claim 1 wherein: themembrane comprises regenerated cellulose or cellulose ester.
 6. Themicrofluidic device of claim 1 wherein: the membrane is hydrophilic. 7.The microfluidic device of claim 1 wherein: the wetting control layercomprises a fluoropolymer.
 8. The microfluidic device of claim 7wherein: the first fluid comprises a fluorinated oil.
 9. Themicrofluidic device of claim 1 wherein: the wetting control layercomprises a polymeric material selected from the group consisting offluoroakylenes and blends and copolymers thereof.
 10. The microfluidicdevice of claim 1 wherein: the wetting control layer comprisesfluorinated ethylene propylene.
 11. The microfluidic device of claim 1wherein: the storage layer includes a fluorophilic coating.
 12. Themicrofluidic device of claim 1 wherein: the reservoir, the membrane, thewetting control layer and the storage layer are reversibly securedtogether by clamping or are laminated together.
 13. The microfluidicdevice of claim 1 wherein: the fluid passageway is aligned with thereservoir.
 14. The microfluidic device of claim 1 wherein: a pluralityof storage wells are positioned within the first fluid path, thereservoir layer defines a plurality of reservoirs, and each reservoir isaligned with one of the storage wells.
 15. The microfluidic device ofclaim 1 wherein: the storage layer comprises polyurethane, and thereservoir layer comprises polydimethylsiloxane.
 16. The microfluidicdevice of claim 1 wherein: the storage layer and the reservoir layereach comprise plastic, fluoroplastic, or glass. 17-119. (canceled) 120.An apparatus for controlling a reaction or a phase transition, theapparatus comprising: the device of claim 1; a source of oil in fluidcommunication with the microfluidic channel; and a source of an aqueousfluid in fluid communication with the reservoir.
 121. The apparatus ofclaim 120 wherein: the membrane is a dialysis membrane.
 122. Theapparatus of claim 120 wherein: hydrostatic pressure regulates transportfluxes across the membrane.
 123. The apparatus of claim 120 furthercomprising: a pressure controller for regulating transport fluxes acrossthe membrane.
 124. The apparatus of claim 120 wherein: the aqueous fluidis a buffer.
 125. The apparatus of claim 120 wherein: outlets of themicrofluidic channel are blocked.
 126. The apparatus of claim 120wherein: an outlet of the reservoir is open.
 127. The apparatus of claim120 further comprising: a microscope for monitoring the reaction or thephase transition in the device.
 128. The apparatus of claim 120 wherein:the reaction is one or more of the following: steady-state andself-assembly reactions at or far from equilibrium; perturbationanalysis of reaction networks; cell synchronization; cell and tissuedifferentiation; and/or chemostat reactions with cells and cellpopulations.
 129. The apparatus of claim 120 wherein: the phasetransition is one or more of the following: crystallization andco-crystallization of small molecules, biological macromolecules,colloids and combinations thereof; liquid crystal phase transitions;gelation; liquid-liquid separation; protein folding; and/or DNA meltingor condensation.
 130. The apparatus of claim 120 wherein: the reactionis a chemostat reaction with cells. 131-134. (canceled)